Molecular Detection of Human Bacterial Pathogens

EDITED BY

DONGYOU LIU

MOLECULAR DETECTION OF HUMAN BACTERIAL PATHOGENS

MOLECULAR DETECTION OF HUMAN BACTERIAL PATHOGENS EDITED BY

DONGYOU LIU

Boca Raton London New York

CRC Press is an imprint of the Taylor & Francis Group, an informa business

CRC Press Taylor & Francis Group 6000 Broken Sound Parkway NW, Suite 300 Boca Raton, FL 33487-2742 © 2011 by Taylor & Francis Group, LLC CRC Press is an imprint of Taylor & Francis Group, an Informa business No claim to original U.S. Government works Version Date: 20110629 International Standard Book Number-13: 978-1-4398-1239-6 (eBook - PDF) This book contains information obtained from authentic and highly regarded sources. Reasonable efforts have been made to publish reliable data and information, but the author and publisher cannot assume responsibility for the validity of all materials or the consequences of their use. The authors and publishers have attempted to trace the copyright holders of all material reproduced in this publication and apologize to copyright holders if permission to publish in this form has not been obtained. If any copyright material has not been acknowledged please write and let us know so we may rectify in any future reprint. Except as permitted under U.S. Copyright Law, no part of this book may be reprinted, reproduced, transmitted, or utilized in any form by any electronic, mechanical, or other means, now known or hereafter invented, including photocopying, microfilming, and recording, or in any information storage or retrieval system, without written permission from the publishers. For permission to photocopy or use material electronically from this work, please access www.copyright.com (http://www.copyright.com/) or contact the Copyright Clearance Center, Inc. (CCC), 222 Rosewood Drive, Danvers, MA 01923, 978-750-8400. CCC is a not-for-profit organization that provides licenses and registration for a variety of users. For organizations that have been granted a photocopy license by the CCC, a separate system of payment has been arranged. Trademark Notice: Product or corporate names may be trademarks or registered trademarks, and are used only for identification and explanation without intent to infringe. Visit the Taylor & Francis Web site at http://www.taylorandfrancis.com and the CRC Press Web site at http://www.crcpress.com

This volume is dedicated to a group of bacteriologists whose insight, knowledge, and expertise have made the all-inclusive coverage of major human bacterial pathogens a reality.

Contents Preface���������������������������������������������������������������������������������������������������������������������������������������������������������������������������������������� xv Editor����������������������������������������������������������������������������������������������������������������������������������������������������������������������������������������xvii Contributors�������������������������������������������������������������������������������������������������������������������������������������������������������������������������������xix Chapter 1 Introductory Remarks��������������������������������������������������������������������������������������������������������������������������������������������� 1 Dongyou Liu

Section I  Actinobacteria Chapter 2 Actinomadura�������������������������������������������������������������������������������������������������������������������������������������������������������� 13 Martha E. Trujillo Chapter 3 Actinomyces����������������������������������������������������������������������������������������������������������������������������������������������������������� 23 Debarati Paul, D. Madhusudan Reddy, Dipalok Mukherjee, Biswajit Paul, and Debosmita Paul Chapter 4 Atopobium������������������������������������������������������������������������������������������������������������������������������������������������������������� 31 Piet Cools, Hans Verstraelen, Mario Vaneechoutte, and Rita Verhelst Chapter 5 Bifidobacterium����������������������������������������������������������������������������������������������������������������������������������������������������� 45 Abelardo Margolles, Patricia Ruas-Madiedo, Clara G. de los Reyes-Gavilán, Borja Sánchez, and Miguel Gueimonde Chapter 6 Corynebacterium�������������������������������������������������������������������������������������������������������������������������������������������������� 59 Luis M. Mateos, Michal Letek, Almudena F. Villadangos, María Fiuza, Efrén Ordoñez, and José A. Gil Chapter 7 Cryptobacterium��������������������������������������������������������������������������������������������������������������������������������������������������� 75 Futoshi Nakazawa Chapter 8 Gardnerella����������������������������������������������������������������������������������������������������������������������������������������������������������� 81 Rita Verhelst, Hans Verstraelen, Piet Cools, Guido Lopes dos Santos Santiago, Marleen Temmerman, and Mario Vaneechoutte Chapter 9 Gordonia��������������������������������������������������������������������������������������������������������������������������������������������������������������� 95 Brent A. Lasker, Benjamin D. Moser, and June Brown Chapter 10 Micrococcus and Kocuria������������������������������������������������������������������������������������������������������������������������������������111 Dongyou Liu Chapter 11 Mycobacterium����������������������������������������������������������������������������������������������������������������������������������������������������� 117 Shubhada Shenai and Camilla Rodrigues vii

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Chapter 12 Nocardia�������������������������������������������������������������������������������������������������������������������������������������������������������������� 127 Veronica Rodriguez-Nava, Frédéric Laurent, and Patrick Boiron Chapter 13 Propionibacterium���������������������������������������������������������������������������������������������������������������������������������������������� 137 Andrew McDowell and Sheila Patrick Chapter 14 Rhodococcus������������������������������������������������������������������������������������������������������������������������������������������������������� 155 Debarati Paul, Dipaloke Mukherjee, Soma Mukherjee, and Debosmita Paul Chapter 15 Saccharopolyspora���������������������������������������������������������������������������������������������������������������������������������������������� 169 Caroline Duchaine and Yvon Cormier Chapter 16 Streptomyces��������������������������������������������������������������������������������������������������������������������������������������������������������175 Angel Manteca, Ana Isabel Pelaez, Maria del Mar Garcia-Suarez, and Francisco J. Mendez Chapter 17 Tropheryma����������������������������������������������������������������������������������������������������������������������������������������������������������181 Shoo Peng Siah, Han Shiong Siah, and Dongyou Liu Chapter 18 Tsukamurella������������������������������������������������������������������������������������������������������������������������������������������������������� 189 Mireille M. Kattar

Section II   Firmicutes and Tenericutes Bacilli Chapter 19 Abiotrophia��������������������������������������������������������������������������������������������������������������������������������������������������������� 201 Marco Arosio and Annibale Raglio Chapter 20 Aerococcus�����������������������������������������������������������������������������������������������������������������������������������������������������������211 Debarati Paul, D. Madhusudan Reddy, and Dipalok Mukherjee Chapter 21 Bacillus�����������������������������������������������������������������������������������������������������������������������������������������������������������������219 Noura Raddadi, Ameur Cherif, and Daniele Daffonchio Chapter 22 Enterococcus������������������������������������������������������������������������������������������������������������������������������������������������������� 231 Teresa Semedo-Lemsaddek, Paula Lopes Alves, Rogério Tenreiro, and Maria Teresa Barreto Crespo Chapter 23 Granulicatella����������������������������������������������������������������������������������������������������������������������������������������������������� 249 Sheng Kai Tung and Tsung Chain Chang Chapter 24 Lactobacillus������������������������������������������������������������������������������������������������������������������������������������������������������� 257 Ester Sánchez and Yolanda Sanz

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Chapter 25 Leuconostoc�������������������������������������������������������������������������������������������������������������������������������������������������������� 269 María Mar Tomás and Germán Bou Chapter 26 Listeria���������������������������������������������������������������������������������������������������������������������������������������������������������������� 279 Dongyou Liu and Ting Zhang Chapter 27 Paenibacillus������������������������������������������������������������������������������������������������������������������������������������������������������� 295 Matthew R. Pincus, Nicholas Cassai, Jie Ouyang, and Sharvari Dalal Chapter 28 Staphylococcus���������������������������������������������������������������������������������������������������������������������������������������������������� 307 Paolo Moroni, Giuliano Pisoni, Paola Cremonesi, and Bianca Castiglioni Chapter 29 Streptococcus������������������������������������������������������������������������������������������������������������������������������������������������������ 323 Judith Jansen and Lothar Rink

Clostridia Chapter 30 Acidaminococcus������������������������������������������������������������������������������������������������������������������������������������������������ 339 Hélène Marchandin and Estelle Jumas-Bilak Chapter 31 Anaerococcus, Parvimonas, and Peptoniphilus������������������������������������������������������������������������������������������������� 349 Dongyou Liu and Frank W. Austin Chapter 32 Catabacter����������������������������������������������������������������������������������������������������������������������������������������������������������� 361 Susanna K.P. Lau and Patrick C.Y. Woo Chapter 33 Clostridium��������������������������������������������������������������������������������������������������������������������������������������������������������� 367 Juan J. Córdoba, Emilio Aranda, María G. Córdoba, María J. Benito, Dongyou Liu, and Mar Rodríguez Chapter 34 Dialister��������������������������������������������������������������������������������������������������������������������������������������������������������������� 381 Ana Paula Vieira Colombo, Renata Martins do Souto, and Andréa Vieira Colombo Chapter 35 Eubacterium�������������������������������������������������������������������������������������������������������������������������������������������������������� 391 Johanna Maukonen and Maria Saarela Chapter 36 Finegoldia����������������������������������������������������������������������������������������������������������������������������������������������������������� 405 A.C.M. Veloo Chapter 37 Mogibacterium�����������������������������������������������������������������������������������������������������������������������������������������������������415 Reginaldo Bruno Gonçalves, Daniel Saito, and Renato Corrêa Viana Casarin Chapter 38 Peptostreptococcus��������������������������������������������������������������������������������������������������������������������������������������������� 423 Eija Könönen and Jari Jalava

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Chapter 39 Tannerella����������������������������������������������������������������������������������������������������������������������������������������������������������� 437 Derren Ready and Lena Ciric Chapter 40 Veillonella����������������������������������������������������������������������������������������������������������������������������������������������������������� 447 Dongyou Liu

Mollicutes Chapter 41 Mycoplasma�������������������������������������������������������������������������������������������������������������������������������������������������������� 455 Rama Chaudhry, Bishwanath Kumar Chourasia, and Anupam Das Chapter 42 Ureaplasma��������������������������������������������������������������������������������������������������������������������������������������������������������� 469 Ken B. Waites, Li Xiao, Vanya Paralanov, and John I. Glass

Section III  Bacteroidetes, Chlamydiae, and Fusobacteria Chapter 43 Bacteroides���������������������������������������������������������������������������������������������������������������������������������������������������������� 491 Rama Chaudhry and Nidhi Sharma Chapter 44 Capnocytophaga������������������������������������������������������������������������������������������������������������������������������������������������� 501 Kazuyuki Ishihara, Satoru Inagaki, and Atsushi Saito Chapter 45 Chlamydia������������������������������������������������������������������������������������������������������������������������������������������������������������511 Jens Kjølseth Møller Chapter 46 Chlamydophila���������������������������������������������������������������������������������������������������������������������������������������������������� 523 Chengming Wang, Bernhard Kaltenboeck, and Konrad Sachse Chapter 47 Elizabethkingia, Chryseobacterium, and Bergeyella����������������������������������������������������������������������������������������� 537 Dongyou Liu Chapter 48 Fusobacterium���������������������������������������������������������������������������������������������������������������������������������������������������� 543 Dongyou Liu and Xiaoming Dong Chapter 49 Leptotrichia and Leptotrichia-Like Organisms�������������������������������������������������������������������������������������������������� 555 Emenike Ribs K. Eribe and Ingar Olsen Chapter 50 Porphyromonas�������������������������������������������������������������������������������������������������������������������������������������������������� 567 Stefan Rupf, Wolfgang Pfister, and Klaus Eschrich Chapter 51 Prevotella������������������������������������������������������������������������������������������������������������������������������������������������������������ 585 Mario J. Avila-Campos, Maria R.L. Simionato, and Elerson Gaetti-Jardim Jr.

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Section IV  Proteobacteria Alphaproteobacteria Chapter 52 Anaplasma����������������������������������������������������������������������������������������������������������������������������������������������������������� 601 Marina E. Eremeeva and Gregory A. Dasch Chapter 53 Bartonella������������������������������������������������������������������������������������������������������������������������������������������������������������� 617 Alessandra Ciervo, Marco Cassone, Fabiola Mancini, and Lorenzo Ciceroni Chapter 54 Brucella��������������������������������������������������������������������������������������������������������������������������������������������������������������� 629 Sascha Al Dahouk, Heinrich Neubauer, and Herbert Tomaso Chapter 55 Ehrlichia�������������������������������������������������������������������������������������������������������������������������������������������������������������� 647 Jere W. McBride, Juan P. Olano, and Nahed Ismail Chapter 56 Ochrobactrum����������������������������������������������������������������������������������������������������������������������������������������������������� 659 Corinne Teyssier and Estelle Jumas-Bilak Chapter 57 Orientia��������������������������������������������������������������������������������������������������������������������������������������������������������������� 671 Harsh Vardhan Batra and Diprabhanu Bakshi Chapter 58 Rickettsia������������������������������������������������������������������������������������������������������������������������������������������������������������� 683 Marina E. Eremeeva and Gregory A. Dasch

Betaproteobacteria Chapter 59 Achromobacter���������������������������������������������������������������������������������������������������������������������������������������������������� 703 Beth Mutai and Yi-Wei Tang Chapter 60 Bordetella������������������������������������������������������������������������������������������������������������������������������������������������������������ 709 Diego Omar Serra, Alejandra Bosch, and Osvaldo Miguel Yantorno Chapter 61 Burkholderia������������������������������������������������������������������������������������������������������������������������������������������������������� 723 Karlene H. Lynch and Jonathan J. Dennis Chapter 62 Eikenella�������������������������������������������������������������������������������������������������������������������������������������������������������������� 737 Javier Enrique Botero, Adriana Jaramillo, and Adolfo Contreras Chapter 63 Kingella��������������������������������������������������������������������������������������������������������������������������������������������������������������� 745 Dongyou Liu

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Chapter 64 Laribacter������������������������������������������������������������������������������������������������������������������������������������������������������������751 Patrick C.Y. Woo and Susanna K.P. Lau Chapter 65 Neisseria�������������������������������������������������������������������������������������������������������������������������������������������������������������� 757 Paola Stefanelli Chapter 66 Ralstonia������������������������������������������������������������������������������������������������������������������������������������������������������������� 769 Michael P. Ryan, J. Tony Pembroke, and Catherine C. Adley

Gammaproteobacteria Chapter 67 Acinetobacter������������������������������������������������������������������������������������������������������������������������������������������������������ 781 Dongyou Liu Chapter 68 Aeromonas����������������������������������������������������������������������������������������������������������������������������������������������������������� 789 Germán Naharro, Pedro Rubio, and José Maria Luengo Chapter 69 Aggregatibacter��������������������������������������������������������������������������������������������������������������������������������������������������� 801 Dongyou Liu Chapter 70 Cardiobacterium��������������������������������������������������������������������������������������������������������������������������������������������������811 Efthimia Petinaki and George N. Dalekos Chapter 71 Cedecea����������������������������������������������������������������������������������������������������������������������������������������������������������������817 Maria Dalamaga and Georgia Vrioni Chapter 72 Citrobacter���������������������������������������������������������������������������������������������������������������������������������������������������������� 827 Ignasi Roca and Jordi Vila Chapter 73 Coxiella��������������������������������������������������������������������������������������������������������������������������������������������������������������� 837 Claire Pelletier Chapter 74 Enterobacter�������������������������������������������������������������������������������������������������������������������������������������������������������� 853 Angelika Lehner, Roger Stephan, Seamus Fanning, and Carol Iversen Chapter 75 Escherichia���������������������������������������������������������������������������������������������������������������������������������������������������������� 869 P. Elizaquível, G. Sánchez, and R. Aznar Chapter 76 Francisella tularensis����������������������������������������������������������������������������������������������������������������������������������������� 881 Kiersten J. Kugeler and Jeannine M. Petersen Chapter 77 Haemophilus������������������������������������������������������������������������������������������������������������������������������������������������������� 893 Dongyou Liu, Jianshun Chen, and Weihuan Fang

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Chapter 78 Klebsiella������������������������������������������������������������������������������������������������������������������������������������������������������������� 905 Beatriz Meurer Moreira, Ana Cristina Gales, Maria Silvana Alves, and Rubens Clayton da Silva Dias Chapter 79 Legionella�������������������������������������������������������������������������������������������������������������������������������������������������������������919 Dianna J. Bopp, Kimberlee A. Musser, and Elizabeth J. Nazarian Chapter 80 Moraxella������������������������������������������������������������������������������������������������������������������������������������������������������������ 929 Suzanne J.C. Verhaegh and John P. Hays Chapter 81 Pasteurella���������������������������������������������������������������������������������������������������������������������������������������������������������� 945 Francis Dziva and Henrik Christensen Chapter 82 Photobacterium��������������������������������������������������������������������������������������������������������������������������������������������������� 959 Carlos R. Osorio and Manuel L. Lemos Chapter 83 Plesiomonas�������������������������������������������������������������������������������������������������������������������������������������������������������� 969 Jesús A. Santos, Andrés Otero, and María-Luisa García-López Chapter 84 Proteus���������������������������������������������������������������������������������������������������������������������������������������������������������������� 981 Antoni Róz˙alski and Paweł Sta˛czek Chapter 85 Providencia��������������������������������������������������������������������������������������������������������������������������������������������������������� 997 Brunella Posteraro, Maurizio Sanguinetti, and Patrizia Posteraro Chapter 86 Pseudomonas���������������������������������������������������������������������������������������������������������������������������������������������������� 1009 Timothy J. Kidd, David M. Whiley, Scott C. Bell, and Keith Grimwood Chapter 87 Salmonella��������������������������������������������������������������������������������������������������������������������������������������������������������� 1023 Mathilde H. Josefsen, Charlotta Löfström, Katharina E.P. Olsen, Kåre Mølbak, and Jeffrey Hoorfar Chapter 88 Serratia�������������������������������������������������������������������������������������������������������������������������������������������������������������� 1037 Kevin B. Laupland and Deirdre L. Church Chapter 89 Shigella�������������������������������������������������������������������������������������������������������������������������������������������������������������� 1049 K.R. Schneider, L.K. Strawn, K.A. Lampel, and B.R. Warren Chapter 90 Stenotrophomonas�������������������������������������������������������������������������������������������������������������������������������������������� 1063 Martina Adamek and Stephan Bathe Chapter 91 Vibrio����������������������������������������������������������������������������������������������������������������������������������������������������������������� 1073 Asim Bej Chapter 92 Yersinia�������������������������������������������������������������������������������������������������������������������������������������������������������������� 1089 Mikael Skurnik, Peter Rådström, Rickard Knutsson, Bo Segerman, Saija Hallanvuo, Susanne Thisted Lambertz, Hannu Korkeala, and Maria Fredriksson-Ahomaa

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Contents

Epsilonproteobacteria Chapter 93 Arcobacter���������������������������������������������������������������������������������������������������������������������������������������������������������� 1111 Kurt Houf Chapter 94 Campylobacter���������������������������������������������������������������������������������������������������������������������������������������������������1125 Rodrigo Alonso, Cecilia Girbau, and Aurora Fernández Astorga Chapter 95 Helicobacter�������������������������������������������������������������������������������������������������������������������������������������������������������1141 Athanasios Makristathis and Alexander M. Hirschl

Section V  Spirochaetes Chapter 96 Borrelia��������������������������������������������������������������������������������������������������������������������������������������������������������������1155 Nataliia Rudenko, Maryna Golovchenko, James H. Oliver Jr., and Libor Grubhoffer Chapter 97 Leptospira����������������������������������������������������������������������������������������������������������������������������������������������������������1169 Rudy A. Hartskeerl Chapter 98 Treponema���������������������������������������������������������������������������������������������������������������������������������������������������������� 1189 Rita Castro and Filomena Martins Pereira

Section VI  Pan-Bacterial Detection Chapter 99 Pan-Bacterial Detection of Sepsis-Causative Pathogens���������������������������������������������������������������������������������� 1203 Roland P.H. Schmitz and Marc Lehmann Chapter 100 Metagenomic Approaches for Bacterial Detection and Identification���������������������������������������������������������������1215 Chaysavanh Manichanh

Preface Bacteria are small, unicellular organisms that are invisible to the naked eye but are nonetheless present ubiquitously and abundantly in all environments. Although a majority of bacteria are free-living and symbiotic, some are capable of leading a parasitic life, inducing a range of disease syndromes in human and animal hosts during the process. Among the most devastating bacterial pathogens, Yersinia pestis, the causative agent of bubonic plague, was responsible for three major human pandemics in history, killing 200 million people prior to the advent of antibiotics. The current, most common fatal bacterial diseases are tuberculosis (caused by Mycobacterium tuberculosis), killing about 2 million people a year alone, and cholera (caused by Vibrio cholerae). Other globally important bacterial diseases include pneumonia (caused by Streptococcus and Pseudomonas), tetanus, typhoid fever, diphtheria, syphilis, and leprosy. Traditionally, bacteria have been identified and diagnosed with the help of various phenotypic procedures, such as Gram stain, morphological, biochemical, and serological examination. Since the phenotypic techniques are often slow and lack desired specificity and reproducibility, nucleic acid amplification technologies such as polymerase chain reaction (PCR) have played an increasingly prominent role in the laboratory diagnosis of bacterial infections. Given their ability to specifically detect a single copy of bacterial nucleic acid template in a matter of hours, PCR-based assays offer unsurpassed sensitivity, specificity, accuracy, precision, and result availability for bacterial identification. The recent advances in instrumentation automation and probe chemistries have facilitated the development of real-time PCR that provides a convenient platform for high throughput detection and quantitation of bacterial pathogens in clinical specimens. Considering that numerous original molecular protocols and subsequent modifications have been described and

scattered in various journals and monographs, it has become difficult if not impossible for someone who has not been directly involved in the development of original or modified protocols to know which are most appropriate to adopt for accurate identification of bacterial pathogens of interest. The purpose of this volume is to address this issue, with international scientists in respective bacterial pathogen research and diagnosis providing expert summaries on current diagnostic approaches for major human bacterial pathogens. Each chapter consists of a brief review of the classification, epidemiology, clinical features, and diagnosis of an important pathogenic bacterial genus; an outline of clinical sample collection and preparation procedures; a selection of representative stepwise molecular protocols; and a discussion on further research requirements relating to improved diagnosis. This book represents a reliable and convenient reference on molecular detection and identification of major human bacterial pathogens; an indispensable tool for upcoming and experienced medical, veterinary, and industrial laboratory scientists engaged in bacterial characterization; and an essential textbook for undergraduate and graduate students majoring in bacteriology. A comprehensive and inclusive book such as this is undoubtedly beyond an individual’s capacity. I am fortunate and honored to have a large panel of bacteriologists as chapter contributors, whose detailed knowledge and technical insights on human bacterial pathogen detection have greatly enriched this book. In addition, the professionalism and dedication of executive editor Barbara Norwitz and senior project coordinator Jill Jurgensen at CRC Press have enhanced its presentation. Finally, without the understanding and support of my family, Liling Ma, Brenda, and Cathy, the compilation of this all-encompassing volume would have not been possible.

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Editor Dongyou Liu, PhD, undertook his veterinary science education at Hunan Agricultural University, China. Upon graduation, he received an Overseas Postgraduate Scholarship from the Chinese Ministry of Education to pursue further training at the University of Melbourne, Australia, where he worked toward improved immunological diagnosis of human hydatid disease. During the past two decades, he has crisscrossed between research and clinical laboratories in Australia and the United States of America, with focuses on molecular characterization and virulence determination of microbial pathogens such as ovine footrot bacterium (Dichelobacter

nodosus), dermatophyte fungi (Trichophyton, Microsporum, and Epidermophyton), and listeriae (Listeria species). He is the senior author of more than 50 original research and review articles in various international journals and the editor of the recently released Handbook of Listeria monocytogenes, Handbook of Nucleic Acid Purification, Molecular Detection of Foodborne Pathogens, and Molecular Detection of Human Viral Pathogens, as well as the forthcoming Molecular Detection of Human Fungal Pathogens and Molecular Detection of Human Parasitic Pathogens, all of which are published by CRC Press.

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Contributors Martina Adamek Karlsruhe Institute of Technology Institute of Functional Interfaces Karlsruhe, Germany

Frank W. Austin College of Veterinary Medicine Mississippi State University Starkville, Mississippi

Dianna J. Bopp Wadsworth Center New York State Department of Health Albany, New York

Catherine C. Adley Microbiology Laboratory Department of Chemical and Environmental Sciences University of Limerick Limerick, Ireland

Mario J. Avila-Campos Department of Microbiology Institute of Biomedical Science University of São Paulo São Paulo, Brazil

Alejandra Bosch Facultad de Ciencias Exactas Centro de Investigación y Desarrollo en Fermentaciones Industriales Universidad Nacional de La Plata La Plata, Argentina

Sascha Al Dahouk Department of Internal Medicine III RWTH Aachen University Aachen, Germany Rodrigo Alonso Faculty of Pharmacy Department of Immunology, Microbiology and Parasitology University of the Basque Country Vitoria-Gasteiz, Spain Maria Silvana Alves Faculdade de Farmácia e Bioquímica Universidade Federal de Juiz de Fora Minas Gerais, Brazil

R. Aznar Department of Microbiology and Ecology University of Valencia Valencia, Spain Diprabhanu Bakshi Syngene International Bangalore, India Stephan Bathe German National Academic Foundation Bonn, Germany Harsh Vardhan Batra Defence R & D Establishment Gwalior, India

Paula Lopes Alves Instituto de Biologia Experimental e Tecnológica Oeiras, Portugal

Asim Bej Department of Biology University of Alabama at Birmingham Birmingham, Alabama

Emilio Aranda Ciencia y Tecnología de los Alimentos Escuela de Ingenierías Agrarias Badajoz, Spain

Scott C. Bell Department of Thoracic Medicine The Prince Charles Hospital Brisbane, Australia

Marco Arosio USC Microbiology and Virology A.O. Ospedali Riuniti Bergamo, Italy

María J. Benito Ciencia y Tecnología de los Alimentos Escuela de Ingenierías Agrarias Badajoz, Spain

Aurora Fernández Astorga Faculty of Pharmacy Department of Immunology, Microbiology and Parasitology University of the Basque Country Vitoria-Gasteiz, Spain

Patrick Boiron Research group on Bacterial Opportunistic Pathogens and Environment Université de Lyon Lyon, France

Javier Enrique Botero School of Dentistry Grupo de Investigación Básica y Clínica en Periodoncia y Oseointegración Universidad de Antioquia Medellín, Colombia Germán Bou Servicio Microbiología Complejo Hospitalario Universitario La Coruña La Coruña, Spain June Brown Actinomycete Reference Laboratory Bacterial Zoonoses Branch Division of Foodborne, Bacterial and Mycotic Diseases National Center for Zoonotic, VectorBorne, and Enteric Diseases Centers for Disease Control and Prevention Atlanta, Georgia Renato Corrêa Viana Casarin Department of Oral Diagnostics Division of Microbiology and Immunology Piracicaba Dental School University of Campinas Campinas, Brazil Nicholas Cassai Department of Pathology and Laboratory Medicine New York Harbor VA Medical Center Brooklyn, New York xix

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Marco Cassone Sbarro Health Research Organization College of Science and Technology Temple University Philadelphia, Pennsylvania Bianca Castiglioni Department of Veterinary Pathology, Hygiene and Public Health University of Milan Milan, Italy Rita Castro Instituto de Higiene e Medicina Tropical Universidade Nova de Lisboa Lisbon, Portugal Tsung Chain Chang Department of Medical Laboratory Science and Biotechnology College of Medicine National Cheng Kung University Tainan, Taiwan Rama Chaudhry Department of Microbiology All India Institute of Medical Sciences New Delhi, India Jianshun Chen Institute of Preventive Veterinary Medicine Zhejiang University Zhejiang, China Ameur Cherif Faculté des Sciences de Tunis Laboratoire Microorganismes et Biomolécules Actives Campus Universitaire Tunis, Tunisie Bishwanath Kumar Chourasia Department of Microbiology All India Institute of Medical Sciences New Delhi, India Henrik Christensen Faculty of Life Sciences Department of Veterinary Pathobiology Center for Applied Bioinformatics Copenhagen University Frederiksberg, Denmark

Contributors

Deirdre L. Church Departments of Pathology, Laboratory Medicine, and Medicine University of Calgary Alberta Health Services and Calgary Laboratory Services Alberta, Canada Lorenzo Ciceroni Center for Research and Evaluation of Immunobiologicals Istituto Superiore di Sanità Rome, Italy Alessandra Ciervo Department of Infectious, Parasitic and Immune-Mediated Diseases Istituto Superiore di Sanità Rome, Italy Lena Ciric Microbial Diseases UCL Eastman Dental Institute London, United Kingdom Ana Paula Vieira Colombo Department of Medical Microbiology Institute of Microbiology Federal University of Rio de Janeiro Rio de Janeiro, Brazil Andréa Vieira Colombo Institute of Biomedical Sciences State University of São Paulo Sao Paulo, Brazil Adolfo Contreras School of Dentistry Grupo de Medicina Periodontal Universidad del Valle Cali, Colombia Piet Cools Department of Clinical Biology, Immunology and Microbiology Ghent University Hospital Ghent, Belgium Juan J. Córdoba Facultad de Veterinaria Higiene de los Alimentos Universidad de Extremadura Cáceres, Spain María G. Córdoba Ciencia y Tecnología de los Alimentos Escuela de Ingenierías Agrarias Badajoz, Spain

Yvon Cormier Research Unit Laval Hospital Centre of Pneumology University of Laval Quebec, Canada Paola Cremonesi Department of Veterinary Pathology, Hygiene and Public Health University of Milan Milan, Italy Maria Teresa Barreto Crespo Instituto de Biologia Experimental e Tecnológica Oeiras, Portugal Daniele Daffonchio Dipartimento di Scienze e Tecnologie Alimentari e Microbiologiche Università degli Studi di Milano Milan, Italy Sharvari Dalal Department of Pathology SUNY Downstate Medical Center Brooklyn, New York Maria Dalamaga Laboratory of Clinical Biochemistry Attikon General University Hospital Medical School, National & Kapodistrian University of Athens Athens, Greece George N. Dalekos Department of Medicine, Medical School University of Thessaly Larissa, Greece Anupam Das Department of Microbiology All India Institute of Medical Sciences New Delhi, India Gregory A. Dasch Rickettsial Zoonoses Branch Division of Vector-Borne Diseases Centers for Disease Control and Prevention Atlanta, Georgia

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Contributors

Jonathan J. Dennis Department of Biological Sciences University of Alberta Alberta, Canada Rubens Clayton da Silva Dias Faculdade de Medicina Universidade Federal do Rio de Janeiro Rio de Janeiro, Brazil Xiaoming Dong Beijing Wantai Biological Pharmacy Enterprise Beijing, China Caroline Duchaine Research Unit Laval Hospital Centre of Pneumology University of Laval Quebec, Canada Francis Dziva Division of Microbiology Institute for Animal Health Berkshire, United Kingdom P. Elizaquível Department of Microbiology and Ecology University of Valencia Valencia, Spain Marina E. Eremeeva Rickettsial Zoonoses Branch Division of Vector-Borne Diseases Centers for Disease Control and Prevention Atlanta, Georgia Emenike Ribs K. Eribe Norwegian Geotechnical Institute Oslo, Norway Klaus Eschrich Institute for Biochemistry School of Medicine University of Leipzig Leipzig, Germany Weihuan Fang Institute of Preventive Veterinary Medicine Zhejiang University Zhejiang, China

Seamus Fanning Centre for Food Safety School of Agriculture, Food Science and Veterinary Medicine Veterinary Sciences Centre University College Dublin Dublin, Ireland

John I. Glass J. Craig Venter Institute Rockville, Maryland

María Fiuza Faculty of Biology Section of Microbiology Department of Molecular Biology University of León León, Spain

Reginaldo Bruno Gonçalves Department of Oral Diagnostics Division of Microbiology and Immunology Piracicaba Dental School University of Campinas Campinas, Brazil

Maria Fredriksson-Ahomaa Ludwig-Maximilian University Munich, Germany Elerson Gaetti-Jardim Jr. Department of Oral Pathology São Paulo State University São Paulo, Brazil Ana Cristina Gales Departamento de Medicina Universidade Federal de São Paulo São Paulo, Brazil María-Luisa García-López Veterinary Faculty Department of Food Hygiene and Food Microbiology University of León León, Spain Maria del Mar Garcia-Suarez Area de Microbiologia Departamento de Biologia Funcional Instituto Universitario de Biotecnologia de Asturias Asturias, Spain José A Gil Faculty of Biology Section of Microbiology Department of Molecular Biology University of León Campus de Vegazana León, Spain Cecilia Girbau Faculty of Pharmacy Department of Immunology, Microbiology and Parasitology University of the Basque Country Vitoria-Gasteiz, Spain

Maryna Golovchenko Faculty of Science University of South Bohemia Ceské Budeˇjovice, Czech Republic

Keith Grimwood Queensland Paediatric Infectious Diseases Laboratory Royal Children’s Hospital Queensland Children’s Medical Research Institute Queensland, Australia Libor Grubhoffer Biology Centre Institute of Parasitology Academy of Sciences of the Czech Republic Ceské Budeˇjovice, Czech Republic Miguel Gueimonde Department of Microbiology and Biochemistry of Dairy Products Instituto de Productos Lácteos de Asturias Consejo Superior de Investigaciones Científicas Asturias, Spain Saija Hallanvuo Finnish Food Safety Authority Evira National Public Health Institute Helsinki, Finland Rudy A. Hartskeerl WHO/FAO/OIE and National Collaborating Centre for Reference and Research on Leptospirosis Department KIT Biomedical Research Royal Tropical Institute Amsterdam, the Netherlands John P. Hays Department of Medical Microbiology and Infectious Diseases Rotterdam, the Netherlands

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Contributors

Alexander M. Hirschl Division of Clinical Microbiology Department of Laboratory Medicine Medical University Vienna Vienna, Austria

Adriana Jaramillo School of Dentistry Grupo de Medicina Periodontal Universidad del Valle Cali, Colombia

Jeffrey Hoorfar National Food Institute Technical University of Denmark Søborg, Denmark

Mathilde H. Josefsen National Food Institute Technical University of Denmark Søborg, Denmark

Kurt Houf Faculty of Veterinary Medicine Department of Veterinary Public Health and Food Safety Ghent University Ghent, Belgium

Estelle Jumas-Bilak Universitaire de Montpellier Hôpital Arnaud de Villeneuve Laboratoire de Bactériologie Montpellier, France

Satoru Inagaki Department of Microbiology Oral Health Science Center Tokyo Dental College Chiba, Japan

Bernhard Kaltenboeck Molecular Diagnostics Laboratory Department of Pathobiology College of Veterinary Medicine Auburn University Auburn, Alabama

Kazuyuki Ishihara Department of Microbiology Oral Health Science Center Tokyo Dental College Chiba, Japan Nahed Ismail Department of Pathology Meharry Medical College Nashville, Tennessee Carol Iversen Centre for Food Safety School of Agriculture Food Science and Veterinary Medicine Veterinary Sciences Centre University College Dublin Dublin, Ireland Jari Jalava Department of Infectious Disease Surveillance and Control National Institute for Health and Welfare Turku, Finland Judith Jansen Medical Faculty Institute of Immunology RWTH Aachen University Aachen, Germany

Mireille M. Kattar Department of Laboratory Medicine and Pathology University of Alberta Hospital Alberta, Canada Timothy J. Kidd Queensland Paediatric Infectious Diseases Laboratory Royal Children’s Hospital Queensland Children’s Medical Research Institute Queensland, Australia Rickard Knutsson Swedish National Veterinary Institute Uppsala, Sweden Eija Könönen University of Turku Institute for Dentistry Turku, Finland Hannu Korkeala Faculty of Veterinary Medicine Department of Food and Environmental Hygiene University of Helsinki Helsinki, Finland

Kiersten J. Kugeler Centers for Disease Control and Prevention National Center for Zoonotic VectorBorne and Enteric Diseases Division of Vector-Borne Infectious Diseases, Bacterial Diseases Branch Fort Collins, Colorado Susanne Thisted Lambertz Research and Development Department National Food Administration Uppsala, Sweden K.A. Lampel Food and Drug Administration Division of Microbiology College Park, Maryland Brent A. Lasker Actinomycete Reference Laboratory Bacterial Zoonoses Branch Division of Foodborne, Bacterial and Mycotic Diseases National Center for Zoonotic, VectorBorne, and Enteric Diseases Centers for Disease Control and Prevention Atlanta, Georgia Susanna K.P. Lau Department of Microbiology The University of Hong Kong Hong Kong Kevin B. Laupland Departments of Medicine, Critical Care Medicine, Community Health Sciences, and Pathology and Laboratory Medicine, and Centre for Antimicrobial Resistance University of Calgary, Alberta Health Services, and Calgary Laboratory Services Alberta, Canada Frédéric Laurent Faculté de Médecine Laennec Lyon, France Marc Lehmann SIRS-Lab GmbH Jena, Germany Angelika Lehner Vetsuisse Faculty Institute for Food Safety and Hygiene University of Zurich Zurich, Switzerland

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Contributors

Manuel L. Lemos Department of Microbiology and Parasitology Institute of Aquaculture University of Santiago de Compostela Santiago de Compostela, Galicia, Spain

Angel Manteca Area de Microbiologia Departamento de Biologia Funcional Instituto Universitario de Biotecnologia de Asturias Universidad de Oviedo Asturias, Spain

Michal Letek Faculty of Biology Section of Microbiology Department of Molecular Biology University of León Campus de Vegazana León, Spain

Hélène Marchandin Faculté de Pharmacie Université Montpellier 1 Laboratoire de Bactériologie-VirologieContrôle Microbiologique Montpellier, France

Dongyou Liu BioSecurity Quality Assurance Program Royal College of Pathologists of Australasia New South Wales, Australia

Abelardo Margolles Department of Microbiology and Biochemistry of Dairy Products Instituto de Productos Lácteos de Asturias Consejo Superior de Investigaciones Científicas Asturias, Spain

Charlotta Löfström National Food Institute Technical University of Denmark Søborg, Denmark José Maria Luengo Department of Molecular Biology University of León León, Spain Karlene H. Lynch Department of Biological Sciences University of Alberta Alberta, Canada Athanasios Makristathis Division of Clinical Microbiology Department of Laboratory Medicine Medical University Vienna Vienna, Austria Fabiola Mancini Department of Infectious, Parasitic and Immune-Mediated Diseases Istituto Superiore di Sanità Rome, Italy Chaysavanh Manichanh Digestive System Research Unit University Hospital Vall d’Hebron Bioinformatics and Genomics Program Center for Genomic Regulation Barcelona, Spain

Luis M. Mateos Faculty of Biology Section of Microbiology Department of Molecular Biology University of León Campus de Vegazana León, Spain Johanna Maukonen VTT Technical Research Centre of Finland Espoo, Finland Jere W. McBride Department of Pathology University of Texas Medical Branch at Galveston Galveston, Texas Andrew McDowell Centre of Infection & Immunity School of Medicine, Dentistry and Biomedical Sciences Queen’s University Belfast, Northern Ireland Francisco J. Mendez Area de Microbiologia Departamento de Biologia Funcional Instituto Universitario de Biotecnologia de Asturias Universidad de Oviedo Asturias, Spain

Kåre Mølbak Statens Serum Institut Copenhagen, Denmark Jens Kjølseth Møller Department of Clinical Microbiology Institute of Regional Health Services Research University of Southern Denmark Odense, Denmark Beatriz Meurer Moreira Instituto de Microbiologia Universidade Federal do Rio de Janeiro Rio de Janeiro, Brazil Paolo Moroni Quality Milk Production Services Cornell University Ithaca, New York Benjamin D. Moser Actinomycete Reference Laboratory Bacterial Zoonoses Branch Division of Foodborne, Bacterial and Mycotic Diseases National Center for Zoonotic, VectorBorne, and Enteric Diseases Centers for Disease Control and Prevention Atlanta, Georgia Dipalok Mukherjee Biological Sciences Mississippi State University Starkville, Mississippi Soma Mukherjee Palli Shikha Bhavan Sriniketan, Birbhum West Bengal, India Kimberlee A. Musser Wadsworth Center New York State Department of Health Albany, New York Beth Mutai Global Emerging Infections Surveillance System U.S. Army Medical Research Unit Kenya Medical Research Institute Narombi, Kenya

xxiv

Germán Naharro Department of Animal Health University of León León, Spain Futoshi Nakazawa Department of Oral Microbiology School of Dentistry, Health Sciences University of Hokkaido Hokkaido, Japan

Contributors

Carlos R. Osorio Department of Microbiology and Parasitology Institute of Aquaculture University of Santiago de Compostela Compostela, Spain Jie Ouyang Department of Pathology SUNY Downstate Medical Center Brooklyn, New York

Elizabeth J. Nazarian Wadsworth Center New York State Department of Health Albany, New York

Vanya Paralanov J. Craig Venter Institute Rockville, Maryland

Heinrich Neubauer Friedrich-Loeffler-Institut Institute of Bacterial Infections and Zoonoses Jena, Germany

Sheila Patrick Centre of Infection & Immunity School of Medicine, Dentistry and Biomedical Sciences Queen’s University Belfast, Northern Ireland

Juan P. Olano Department of Pathology University of Texas Medical Branch at Galveston Galveston, Texas

Biswajit Paul Escorts Heart Institute and Research Centre New Delhi, India

James H. Oliver, Jr. Georgia Southern University The James H. Oliver, Jr. Institute of Arthropodology and Parasitology Statesboro, Georgia Ingar Olsen Faculty of Dentistry Institute of Oral Biology University of Oslo Oslo, Norway Katharina E.P. Olsen Statens Serum Institut Copenhagen, Denmark

Debarati Paul College of Veterinary Medicine Mississippi State University Starkville, Mississippi Debosmita Paul Jamia Millia Islamia New Delhi, India Ana Isabel Pelaez Area de Microbiologia Departamento de Biologia Funcional Instituto Universitario de Biotecnologia de Asturias Universidad de Oviedo Asturias, Spain

Efrén Ordoñez Faculty of Biology Section of Microbiology Department of Molecular Biology University of León León, Spain

Claire Pelletier Laboratoire Départemental d’Analyses de Saône-et-Loire Mâcon, France

Andrés Otero Veterinary Faculty Department of Food Hygiene and Food Microbiology University of León León, Spain

J. Tony Pembroke Molecular and Structural Biochemistry Laboratory Department of Chemical and Environmental Sciences University of Limerick Limerick, Ireland

Filomena Martins Pereira Instituto de Higiene e Medicina Tropical Universidade Nova de Lisboa Lisbon, Portugal Jeannine M. Petersen Centers for Disease Control and Prevention National Center for Zoonotic, VectorBorne and Enteric Diseases Division of Vector-Borne Infectious Diseases, Bacterial Diseases Branch Fort Collins, Colorado Efthimia Petinaki Department of Microbiology University of Thessaly Medical School Larissa, Greece Wolfgang Pfister Institute for Medical Microbiology University Hospital of Jena Jena, Germany Matthew R. Pincus Department of Pathology SUNY Downstate Medical Center Brooklyn, New York Giuliano Pisoni Department of Veterinary Pathology, Hygiene and Public Health University of Milan Milan, Italy Brunella Posteraro Institute of Microbiology Università Cattolica del Sacro Cuore Rome, Italy Patrizia Posteraro Laboratory of Clinical Pathology and Microbiology Ospedale San Carlo Rome, Italy Noura Raddadi Dipartimento di Scienze e Tecnologie Alimentari e Microbiologiche Università degli Studi di Milano Milan, Italy Peter Rådström Department of Applied Microbiology Lund University Lund, Sweden

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Contributors

Annibale Raglio USC Microbiology and Virology A.O. Ospedali Riuniti Bergamo, Italy

Patricia Ruas-Madiedo Department of Microbiology and Biochemistry of Dairy Products Instituto de Productos Lácteos de Asturias Consejo Superior de Investigaciones Científicas Asturias, Spain

Borja Sánchez Department of Microbiology and Biochemistry of Dairy Products Instituto de Productos Lácteos de Asturias Consejo Superior de Investigaciones Científicas Asturias, Spain

D. Madhusudan Reddy Palamuru University Andhra Pradesh, India

Pedro Rubio Department of Animal Health University of León León, Spain

Clara G. de los Reyes-Gavilán Department of Microbiology and Biochemistry of Dairy Products Instituto de Productos Lácteos de Asturias Consejo Superior de Investigaciones Científicas Asturias, Spain

Nataliia Rudenko Biology Centre Institute of Parasitology Academy of Sciences of the Czech Republic Cˇ eské Budeˇjovice, Czech Republic

Ester Sánchez Microbial Ecophysiology and Nutrition Research Group Institute of Agrochemistry and Food Technology Spanish Council for Scientific Research Valencia, Spain

Lothar Rink Institute of Immunology Medical Faculty RWTH Aachen University Aachen, Germany

Stefan Rupf Clinic of Operative Dentistry Periodontology and Preventive Dentistry Saarland University Hospital Homburg/Saar, Germany

Ignasi Roca Department of Clinical Microbiology School of Medicine University of Barcelona Barcelona, Spain

Michael P. Ryan Microbiology Laboratory Department of Chemical and Environmental Sciences University of Limerick Limerick, Ireland

Derren Ready Eastman Dental Hospital UCLH NHS Foundation Trust London, United Kingdom

Camilla Rodrigues P D Hinduja National Hospital & Medical Research Centre Mumbai, India Mar Rodríguez Facultad de Veterinaria Higiene de los Alimentos Universidad de Extremadura Cáceres, Spain Veronica Rodriguez-Nava Research group on “Bacterial Opportunistic Pathogens and Environment” Université de Lyon Lyon, France Antoni Róz·alski Department of Immunobiology of Bacteria Institute of Microbiology and Immunology University of Łódz´ Łódz´, Poland

Maria Saarela VTT Technical Research Centre of Finland Espoo, Finland Konrad Sachse Friedrich-Loeffler-Institut (Federal Research Institute for Animal Health) Institute of Molecular Pathogenesis Jena, Germany Atsushi Saito Oral Health Science Center Tokyo Dental College Chiba, Japan Daniel Saito Department of Oral Diagnostics Division of Microbiology and Immunology Piracicaba Dental School University of Campinas Campinas, Brazil

G. Sánchez Department of Biotecnology Institute of Agrochemistry and Food Technology Valencia, Spain Maurizio Sanguinetti Institute of Microbiology Università Cattolica del Sacro Cuore Rome, Italy Guido Lopes dos Santos Santiago Department of Clinical Biology, Immunology and Microbiology Ghent University Hospital Ghent, Belgium Jesús A. Santos Veterinary Faculty Department of Food Hygiene and Food Microbiology University of León León, Spain Yolanda Sanz Microbial Ecophysiology and Nutrition Research Group Institute of Agrochemistry and Food Technology Spanish Council for Scientific Research Valencia, Spain Roland P.H. Schmitz SIRS-Lab GmbH Jena, Germany

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Contributors

K.R. Schneider Food Science and Human Nutrition Department University of Florida Gainesville, Florida

Renata Martins do Souto Department of Medical Microbiology Institute of Microbiology Federal University of Rio de Janeiro Rio de Janeiro, Brazil

María Mar Tomás Servicio Microbiología Complejo Hospitalario Universitario La Coruña La Coruña, Spain

Bo Segerman Swedish National Veterinary Institute Uppsala, Sweden

Paweł Sta˛czek Department of Immunobiology of Bacteria Institute of Microbiology and Immunology University of Łódz´ Łódz´, Poland

Herbert Tomaso Friedrich-Loeffler-Institut Institute of Bacterial Infections and Zoonoses Jena, Germany

Teresa Semedo-Lemsaddek Faculdade de Ciências Universidade de Lisboa Center for Biodiversity Functional and Integrative Genomics Edifício ICAT, Campus da FCUL Lisbon, Portugal Diego Omar Serra Facultad de Ciencias Exactas Centro de Investigación y Desarrollo en Fermentaciones Industriales Universidad Nacional de La Plata La Plata, Argentina Nidhi Sharma Department of Microbiology All India Institute of Medical Sciences New Delhi, India Shubhada Shenai P D Hinduja National Hospital & Medical Research Centre Mumbai, India Han Shiong Siah TPP, Lambells Lagoon Northern Territory, Australia Shoo Peng Siah Human Genetic Signatures New South Wales, Australia Maria R.L. Simionato Department of Microbiology Institute of Biomedical Science University of São Paulo São Paulo, Brazil Mikael Skurnik University of Helsinki and Helsinki University Central Hospital Laboratory Diagnostics Helsinki, Finland

Paola Stefanelli Department of Infectious, Parasitic and Immune-Mediated Diseases Istituto Superiore di Sanità Rome, Italy Roger Stephan Vetsuisse Faculty Institute for Food Safety and Hygiene University of Zurich Zurich, Switzerland L.K. Strawn Department of Food Science Cornell University Ithaca, New York Yi-Wei Tang Departments of Pathology and Medicine Vanderbilt University School of Medicine Nashville, Tennessee Marleen Temmerman Department of Obstetrics and Gynaecology Ghent University Ghent, Belgium Rogério Tenreiro Faculdade de Ciências Universidade de Lisboa Center for Biodiversity Functional and Integrative Genomics Edifício ICAT, Campus da FCUL Lisbon, Portugal Corinne Teyssier Faculté de Pharmacie, BP 14491 Université Montpellier 1 Montpellier, France

Martha E. Trujillo Departmento de Microbiología y Genética Edificio Departamental Universidad de Salamanca Salamanca, Spain Sheng Kai Tung Department of Medical Laboratory Science and Biotechnology College of Medicine National Cheng Kung University Taiwan Mario Vaneechoutte Department of Clinical Biology, Immunology and Microbiology Ghent University Hospital Ghent, Belgium A.C.M. Veloo Department of Medical Microbiology University Medical Center Groningen University of Groningen Groningen, the Netherlands Suzanne J.C. Verhaegh Department of Medical Microbiology and Infectious Diseases, Erasmus MC Rotterdam, the Netherlands Rita Verhelst Department of Clinical Biology, Immunology and Microbiology Ghent University Hospital Ghent, Belgium Hans Verstraelen Department of Obstetrics and Gynaecology Ghent University Hospital Ghent, Belgium

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Contributors

Jordi Vila Department of Clinical Microbiology Hospital Clinic University of Barcelona School of Medicine Barcelona, Spain

Chengming Wang Molecular Diagnostics Laboratory Department of Pathobiology College of Veterinary Medicine Auburn University Auburn, Alabama

Almudena F. Villadangos Section of Microbiology Department of Molecular Biology Faculty of Biology University of León León, Spain

B.R. Warren Research, Quality and   Innovation ConAgra Foods, Inc. Omaha, Nebraska

Georgia Vrioni Department of Microbiology National & Kapodistrian University of Athens Medical School Athens, Greece Ken B. Waites Department of Pathology University of Alabama at Birmingham Birmingham, Alabama

David M. Whiley Queensland Paediatric Infectious Diseases Laboratory Royal Children’s Hospital Queensland Children’s Medical Research Institute Queensland, Australia Patrick C.Y. Woo Department of Microbiology The University of Hong Kong Hong Kong

Li Xiao Department of Pathology University of Alabama at Birmingham Birmingham, Alabama Osvaldo Miguel Yantorno Facultad de Ciencias Exactas Centro de Investigación y Desarrollo en Fermentaciones Industriales Universidad Nacional de La Plata La Plata, Argentina Ting Zhang College of Veterinary Medicine Mississippi State University Starkville, Mississippi

1 Introductory Remarks Dongyou Liu CONTENTS 1.1 Preamble............................................................................................................................................................................... 1 1.2 Bacterial Characteristics....................................................................................................................................................... 2 1.2.1 Classification............................................................................................................................................................. 2 1.2.2 Morphology.............................................................................................................................................................. 2 1.2.3 Biology...................................................................................................................................................................... 4 1.2.4 Genetics.................................................................................................................................................................... 5 1.2.5 Ecological and Medical Importance......................................................................................................................... 5 1.3 Bacterial Identification......................................................................................................................................................... 6 1.3.1 Current Diagnostic Approaches............................................................................................................................... 6 1.3.2 Performance Parameters........................................................................................................................................... 7 1.3.3 Result Interpretation................................................................................................................................................. 7 1.3.4 Standardization, Quality Control, and Assurance.................................................................................................... 8 1.4 Conclusion............................................................................................................................................................................ 9 References...................................................................................................................................................................................... 9

1.1  PREAMBLE Bacteria (singular, bacterium) are small unicellular organisms that are classified taxonomically in the domain Bacteria (or Eubacteria), the kingdom Prokaryotae (or Prokaryota or Monera). The only other domain in the kingdom Prokaryotae covers Archaea (or Archaeobacteria for “ancient bacteria”). With sizes ranging from 10 −7 to 10 −4 mm, prokaryotes are bigger than viruses (10 −8–10 −6 mm), but smaller than eukaryotes (10 −5–103 mm). While both bacteria and archaea may have evolved independently from an ancient common ancestor, eukaryotes may have arisen from ancient bacteria entering into endosymbiotic associations with the ancestors of eukaryotic cells (possibly related to the archaea) to form either mitochondria or hydrogenosomes. A subsequent independent engulfment by some mitochondria-containing eukaryotes of cyanobacterial-like organisms may have led to the formation of chloroplasts in algae and plants. In contrast to the organisms in the eukaryotic kingdoms Protista, Fungi, Plantae, and Animalia, those in the kingdom Prokaryotae lack nuclear membrane (with their DNA usually in a loop or coil), contain few independent membrane-bounded cytoplasmic organelles (e.g., vacuole, endoplasmic reticulum, Golgi apparatus, and mitochondria) apart from chromosome and ribosome, have no unique structures in their plasma membrane and cell wall, and do not undergo endocystosis and exocytosis. In other words, whereas eukaryotic chromosome resides within a membrane-delineated nucleus, bacterial chromosome is located inside the

bacterial cytoplasm. This entails that all cellular events (e.g., translational and transcriptional processes, and interaction of chromosome with other cytoplasmic structures) in prokaryotes occur in the same compartment. Furthermore, while eukaryotic chromosome is packed with histones to form linear chromatin, bacterial chromosome assumes a highly compact supercoiled structure in circular form (and rarely in linear form). Although archaea are similar to bacteria in most aspects of cell structure and metabolism, they differ from bacteria in that being extremophiles, they can live in extreme environments where no other life forms exist. This may be due to the unique structure in archaeal lipids in which the stereochemistry of the glycerol is the reverse of that found in bacteria and eukaryotes, possibly the result of an adaptation on the part of archaea to hyperthermophily. In addition, the archaeal cell wall does not contain muramic acid, which is commonly present in bacteria. The archaeal RNA polymerase core is composed of ten subunits in comparison with four subunits in bacteria. Besides possessing distinct tRNA and rRNA genes, archaea uses eukaryotic-like initiation and elongation factors in protein translation, and their transcription involves TATA-binding proteins and TFIIB as in eukaryotes. Bacteria are ubiquitously distributed in virtually every habitat on earth, and are abundantly present in soil, fresh water, plants, and animals. With an estimated number of 5  nonillion (5 × 1030), bacteria form much of the world’s biomass. Bacteria play an essential role in chemical cycles, 1

2

Molecular Detection of Human Bacterial Pathogens

environmental maintenance, food production, and human wellbeing. However, some bacteria are pathogenic and capable of causing infectious diseases in humans, animals, and plants. Cholera, syphilis, anthrax, leprosy, bubonic plague, and tuberculosis are some of the examples of the deadly human diseases that are attributable to bacteria. Correct identification and detection of bacterial pathogens is not only fundamental to the study of these microorganisms but also critical to the control and prevention of the diseases they cause.

1.2  BACTERIAL CHARACTERISTICS

microbia, Thermotogae, and Verrcomicrobia), whereas  the domain Archaea (Archaeobacteria) is separated into two phyla (Crenarchaeota and Euryarchaeota). Among the 26 phyla in the domain Bacteria, Proteobacteria, and Firmicutes contain the largest numbers of genera and species followed by Cyanobacteria, Bacteroidetes, Spirochaetes, and Flavobacteria. Bacteria from other phyla are comparatively rare, from which fewer genera and species have been described. Most of human pathogenic bacteria are found in the phyla Actinobacteria, Bacteroidetes, Chlamydiae, Firmicutes, Fusobacteria, Proteobacteria, Spirochaetes, and Tenericutes (Table 1.1).

1.2.1  Classification

1.2.2  Morphology

Bacteria are classified on the basis of their differences in morphology (e.g., rod, cocci, spirilla, and filament), cell  wall structure (e.g., gram-negative and gram-positive), growth characteristics (e.g., aerobic and anaerobic), biochemical properties (e.g., fatty acids), and genetic features (e.g., 16S and 23S rRNA). Currently, the domain Bacteria (Eubacteria) is divided into 26 phyla (Acidobacteria, Actinobacteria, Aquificae, Bacteroidetes, Chlamydiae, Chlorobi, Chloroflexi, Chrysiogenetes, Cyanobacteria, Deferribacteres, Deinococcus-Thermus, Dictyoglomi, Fibro­ bacteres, Firmicutes, Fusobacteria, Gemmatimonadetes, Lentisphaerae, Nitrospira, Planctomycetes, Proteobacteria, Spirochaetes, Tenericutes, Thermodesulfobacteria, Thermo­

Bacteria usually measure from 0.2 to 2.0 μm in width and 2–8  μm in length, and are 10 times smaller than eukaryotic cells. On one extreme, a few bacterial species (e.g., Thiomargarita namibiensis, and Epulopiscium fishelsoni) measure up to half a mm long and are visible to the naked eye. On the other extreme, the smallest bacteria in the genus Mycoplasma are only 0.3 μm in size, which are as small as the largest viruses. Bacteria typically assume four distinctive forms: rodlike bacilli, spherical cocci, spiral bacteria (also called spirilla), and filamentous bacteria. Occasionally, a small number of bacterial species may appear tetrahedral or cuboidal in shape. While many bacterial species exist as

TABLE 1.1 Classification and Characteristics of Major Human Bacterial Pathogens Phylum

Class

Actinobacteria

Actinobacteria

Firmicutes

Bacilli Clostridia

Tenericutes

Mollicutes

Bacteroidetes

Bacteroidia

Chlamydiae Fusobacteria Proteobacteria

Flavobacteria Chlamydiae Fusobacteria Alphaproteobacteria Betaproteobacteria Gammaproteobacteria

Epsilonproteobacteria Spirochaetes

Spirochaetes

Brief Description Gram-positive bacteria with high G + C ratio; classification may be assisted through analysis of ferric uptake regulator (fur) and glutamine synthetase Gram-positive cocci or rods with low G + C ratio; presence of cell wall Gram-positive cocci or rods with low G + C ratio; presence of cell wall; some species (e.g., Veillonella) are gram-negative Small bacteria (0.2–0.3 μm in size) with low G + C ratio; absence of cell wall (outer membrane) Gram-negative, anaerobic bacteria; opportunistic pathogens Gram-negative, anaerobic bacteria; opportunistic pathogens Gram-negative bacteria; obligate intracellular pathogens Gram-negative, filamentous, anaerobic bacteria Gram-negative, phototrophic bacteria, with symbiotic properties Gram-negative, aerobic or facultative bacteria some of which are chemolithotrophs, or phototrophs Gram-negative, facultatively or obligately anaerobic bacteria, some of which are highly pathogenic Gram-negative, curved to spirilloid bacteria, inhabiting digestive tract Gram-negative, long, helically coiled (spiral-shaped), chemoheterotrophic, anaerobic bacteria, with length-wise flagella

Notable Human Pathogens Actinomyces, Corynebacterium, Mycobacterium, Nocardia Bacillus, Enterococcus, Listeria, Staphylococcus, Streptococcus Clostridium, Eubacterium, Peptostreptococcus Mycoplasma, Ureaplasma Bacteroides, Porphyromonas, Prevotella Elizabethkingia, Flavobacterium Chlamydia, Chlamydophila Fusobacterium, Leptotrichia Bartonella, Brucella Bordetella, Burkholderia, Neisseria Aeromonas, Klebsiella, Pseudomonas, Salmonella, Vibrio, Yersinia Arcobacter, Campylobacter, Helicobacter Borrelia, Leptospira, Treponema

Introductory Remarks

single cells, others present characteristic patterns such as diploids (pairs), chains, and clusters (“bunch of grapes”). In addition, some bacteria may be elongated to form filaments, which are often surrounded by a sheath containing many individual cells. The elaborated, branched filaments formed by Nocardia may even resemble fungal mycelia in appearance. Frequently, bacteria use quorum sensing to detect surrounding cells, migrate toward each other, and attach to solid surfaces to form dense aggregations called biofilms (bacterial mats, or fruiting bodies), which may measure a few micrometers in thickness to up to half a meter in depth, and which comprise multiple species of bacteria, archaea, and protists (numbering approximately 100,000 cells). The formation of biofilms protects bacteria from host defense mechanisms and antibiotic therapy, contributing to chronic bacterial infections and infections relating to implanted medical devices. Structurally, a bacterial cell is surrounded by a rigid layer (cell wall) that is located externally to the lipid membrane. The cell wall provides structural support and protection, and acts as a filtering mechanism. In addition to prokaryotae, fungi and plantae also possess a cell wall, but animalia and most protista do not. While the bacterial cell wall is made up of peptidoglycan (also called murein, which in turn is composed of polysaccharide chain cross-linked by peptides containing d-amino acids), the archaeal cell wall consists of surface layer proteins (also known as S-layer), pseudopeptidoglycan (pseudomurein), and polysaccharides. By contrast, the fungal cell wall includes chitin, the algal cell wall has glycoprotein and polysaccharides, and the plant cell wall often incorporates cellulose and proteins such as extensins. Based on the ability of bacterial cell wall to retain Gram stain (consisting of crystal violet as primary stain and Gram’s iodine and basic fuchsin as subsequent stain), bacteria are divided into gram-positive and gram-negative categories. The gram-positive bacterial cell wall is composed of several layers of peptidoglycan (which is responsible for retaining the crystal violet dyes during the Gram staining procedure, leading to its purple color) surrounded by a second lipid membrane containing lipopolysaccharides and lipoproteins. Located outside of cytoplasmic membrane, peptidoglycan is a large polymer (formed by poly-N-acetylglucosamine and N-acetylemuramic acid) that contributes to the structural integrity of the bacterial cell wall in addition to countering the osmotic pressure of the cytoplasm. Peptidoglycan is predominant in the cell walls of high and low percentage G + C gram-positive organisms (e.g., actinobacteria and firmicutes). Also imbedded in the gram-positive cell wall are teichoic acids, some of which are lipid linked to form lipoteichoic acids. On the other hand, the gram-negative cell wall has a thin peptidoglycan layer adjacent to the cytoplasmic membrane that contributes to its inability to retain the crystal violet stain upon decolonization with ethanol during the Gram staining procedure (leading to its red or pink color after restaining with basic fuchsin). Apart from the thin peptidoglycan layer, the gram-negative cell wall also

3

has an outer membrane that is formed by phospholipids and lipopolysaccharides. Within the gram-positive bacterial category, there is another distinct group of bacteria (i.e., acid-fast bacteria such as Mycobacterium and Nocardia) that can resist decolorization with an acid-alcohol mixture during the acid-fast (or Ziehl–Neelsen) staining procedure and retain the initial dye carbol fuchsin and appear red. The acid-fast cell wall of Mycobacterium includes a large amount of glycolipids, especially mycolic acids that make up approximately 60% of the acid-fast cell wall in addition to a thin, inner-layer peptidoglycan. The presence of the mycolic acids and other glycolipids impede the entry of chemicals, causing the organisms to grow slowly and be more resistant to chemical agents and lysosomal components of phagocytes than most bacteria. Whereas a vast majority of bacteria possess the gramnegative cell wall, the firmicutes and actinobacteria (previously known as the low percentage G + C and high percentage G + C gram-positive bacteria, respectively) have the gram-positive structure, and the tenericutes (e.g., the genus Mycoplasma) are devoid of a cell wall in spite of their similarity in G + C ratio to the firmicutes. The differences in the cell wall often determine the susceptibility and resistance of bacteria to antibiotics and other therapeutic reagents. Given that Mycoplasma species lack a cell wall, they are unaffected by such commonly used antibiotics such as penicillin and streptomycin that target cell wall synthesis. With their small size (0.3 μm), Mycoplasma species are often identified as a source of contaminating infection in the cell culture (where penicillin and streptomycin are incorporated in the culture media), causing retarded growth of cultured cell lines. The cell wall of bacteria forms part of pathogen-associated molecular patterns (or PAMPs), which are recognized by pattern-recognition receptors (or PRRs) in mammalian hosts to initiate and promote innate and adaptive immune defenses against invading bacteria. Several recognizable extracellular structures are present in bacteria. These include flagella, pili, and fimbriae, which protrude from bacterial cell wall and are involved in bacterial twitching movement as well as interaction with one another and other organisms. Bacterial flagellum (measuring 20 nm in diameter and up to 20 μm in length) is a long, whip-like, and helical projection made up of repeating flagellin protein. The numbers and arrangements of flagella vary among bacterial genera and species. Monotrichous bacteria have a single flagellum, amphitrichous bacteria contain a single flagellum on each of cell poles, lophotrichous bacteria include multiple flagella that are located at one cell pole, and peritrichous bacteria have multiple flagella that are situated at several locations. Flagella in bacteria are powered by a flow of H+ ions (sometimes Na+ ions), and those in archaea are powered by adenosine 5'-triphoshate (ATP). Despite having a similar appearance, eukaryotic flagella (called cilia or undulipodia) differ from prokaryotic flagella in both structure and evolutionary origin. A eukaryotic flagellum is a bundle of nine fused pairs of microtubule doublets surrounding two single microtubules. Eukaryotic flagella are often arranged

4

en masse at the surface of a stationary cell anchored within an organ, lashing back and forth and serving to move fluids along mucous membranes such as trachea. In addition, some eukaryotic cells (e.g., rod photoreceptor cells of eye, olfactory receptor cells of nose, and kinocilium in cochlea of ear) have immotile flagella that function as sensation and signal transduction devises. Pilus and fimbria are proteinaceous, hair- or thread-like appendages in bacteria (particularly of gram-negative category) that are much shorter and thinner than flagellum. Bacteria have up to ten pili (typically 6–7 nm in diameter) whose main function is to connect the bacterium to another of the same or a different species to enable transfer of plasmids between the bacteria (i.e., conjugation). A fimbria (measuring 2–10 nm in diameter and up to several μm in length) is shorter than pilus. A bacterium possesses as many as 1000 fimbriae, which are deployed to attach to surface of another bacterium (to form a biofilm) or host cell (to facilitate invasion). Many pilin proteins are characteristic among bacterial species and subgroups, which have been exploited as targets for serological typing of bacteria (serotypes or serovars). Many bacteria produce capsules or slime layers around their cells, which can protect cells from engulfment by eukaryotic cells (e.g., macrophages), act as antigens for cell recognition, and aid attachment to surfaces and the formation of biofilms. In addition, some gram-positive bacteria (e.g., Bacillus, Clostridium, and Anaerobacter) can form highly resistant, dormant structures called endospores, which contain a central core of cytoplasm with DNA and ribosomes surrounded by a cortex layer and protected by an impermeable and rigid coat. Endospores can survive extreme physical and chemical stresses (e.g., UV lights, γ-radiation, detergents and disinfectants, heat, pressure, and desiccation), and may remain viable for millions of years. Endospore-forming bacteria (e.g., Bacillus anthrax and Clostridium tetanus) are also capable of causing disease. Underneath the lipid membrane is the cytoplasm, which is composed of nutrients (or nutrient storage granules such as glycogen, polyphosphate, sulfur, or polyhydroxyalkanoates), proteins, and other essential components. There is a notable absence of membrane-bound organelles (with the exception of chromosome and ribosome) in the bacterial cytoplasm, although certain subcellular compartments (prokaryotic cytoskeleton), such as carboxysome-containing polyhedral protein shells, have been detected. These polyhedral organelles compartmentalize bacterial metabolism, similar to the function performed by the membrane-bound organelles in eukaryotes. The bacterial chromosome consists of a single circular DNA molecule that is situated together with associated proteins and RNA in an irregularly shaped body called the nucleoid. The bacterial ribosomes are responsible for production of proteins.

1.2.3  Biology Bacteria utilize many metabolic pathways (e.g., glycolysis, electron transport chains, chemiosmosis, cellular respiration,

Molecular Detection of Human Bacterial Pathogens

and photosynthesis) and thus virtually all carbon or energy supplies for their maintenance and growth. They are easily grown using either solid or liquid media (e.g., Luria Bertani broth). Solid growth media (e.g., agar plates) are useful for isolation of pure cultures of a bacterial strain, and liquid growth media are employed to generate bulk quantities of bacterial cells. In addition, selective media (containing specific nutrients and antibiotics) assist the isolation and identification of specific bacterial organisms. As single-celled organisms, prokaryotes reproduce by asexual binary fission, which begins with DNA replication within the cell until the entire prokaryotic DNA is duplicated. The two chromosomes then separate as the cell grows and the cell membrane invaginates, splitting the cell into two daughter cells. This reproductive process is highly efficient and leads to exponential growth of bacteria. In fact, under optimal growth conditions, Escherichia coli cells can double every 20 min. Because bacteria are able to multiply rapidly with minimal nutritional requirements, they are abundant in virtually every habitat on earth. In soil, bacteria live by degrading organic compounds and assist in soil formation. In aquatic environments such as ponds, streams, lakes, rivers, seas, and oceans, bacteria such as cyanobacteria (sometimes called blue-green algae because of their color) utilize their chlorophylls to capture energy from the sunlight. In the depths of the sea, bacteria obtain energy from oxidizing or reducing naturally occurring sulfur compounds. In humans and animals, bacteria are found in large numbers on the skin, the respiratory and digestive tracts, and other parts of the body, constituting a normal microbiota in an essentially symbiotic relationship with mutual benefits. Although the vast majority of bacteria are harmless and sometimes even beneficial to their hosts, a few have the capacity to take advantage of temporary weakness in the host (e.g., injury and/or impaired immune function) to cause diseases of varying severity. In a high-nutrient environment, the growth cycle of bacteria usually undergoes three phases. The first phase (the lag phase) is a period of slow growth with the bacterial cells adapting to the high-nutrient environment and preparing for fast growth. In the lag phase, the cell replicates its DNA and makes all the other molecules (e.g., ribosomes, membrane transport proteins) needed for the new cell. The second phase (the logarithmic phase or “log” phase, also known as the exponential phase) occurs when DNA replication stops, and is characterized by rapid cell division and exponential growth. The rate at which cells grow during this phase is known as the growth rate, and the time it takes the cells to double is known as the generation time. During the log phase, nutrients are metabolized at maximum speed until one of the nutrients is depleted, which poses a negative impact on growth. The final phase (the stationary phase) results from the depletion of nutrients. During the stationary phase, the cells decrease their metabolic activity and consume nonessential cellular proteins. As a transition from rapid growth to a stress response state, there is heightened expression of genes involved in DNA repair, antioxidant metabolism,

5

Introductory Remarks

and nutrient transport. Although the entire cycle of bacterial growth takes about an hour, a rapidly growing bacterial cell carries out multiple rounds of replication simultaneously, which helps to shorten the doubling time for most bacteria to about 20 min.

1.2.4  Genetics Bacteria have a single circular chromosome that ranges in size from only 160,000 bp (base pairs) (e.g., Candidatus Carsonella ruddii) to 12,200,000 bp (e.g., Sorangium cellulosum). However, Borrelia burgdorferi, the causal agent for Lyme disease, contains a single linear chromosome. In addition, bacteria may possess small extrachromosomal DNA called plasmid, which ranges from 1 to 400 kb in size and comprises genes or gene cassettes for antibiotic resistance or virulence factors. As plasmids have at least an origin of replication (or ori)—a starting point for DNA replication— they are capable of autonomous replication independent of the chromosomal DNA. A plasmid that integrates into the chromosomal DNA is called episome, which permits its duplication with every cell division of the host. Some viruses (bacteriophages or phages) may also exist in bacteria, with some merely infecting and lysing their host bacteria, while others inserting into the bacterial chromosome. Phages are usually made up of a nucleic acid core (e.g., ssRNA, dsRNA, ssDNA, or dsDNA measuring 5–500 bp in length) with an outer protein hull. A phage containing particular genes may contribute to its host’s phenotype, as illustrated by the evolution of Escherichia coli O157:H7 and Clostridium botulinum, which are converted from harmless ancestral bacteria into lethal pathogens through the integration of phages harboring toxin genes. Being the key component of the ribosome, ribosomal RNA molecules (rRNA) consists of two complex folded subunits of differing sizes (small and large), whose main functions are to provide a mechanism for decoding messenger RNA (mRNA) into amino acids (at center of small ribosomal subunit) and to interact with transfer RNA (tRNA) during translation by providing petidyltransferase activity (large subunit). Whereas the two rRNA subunits in eukaryotes have sedimentation coefficiency values of 40S (Svedberg units) and 60S, those in bacteria measure 30S and 50S, respectively. In virtually all organisms, the small rRNA subunit (40S in eukaryotes and 30S in bacteria) contains a single RNA species (i.e., 18S rRNA in eukaryotes and 16S rRNA in bacteria); the large rRNA subunit (60S) in eukaryotes comprises three RNA species (5S, 5.8S, and 25/28S rRNA), while that (50S) in bacteria contains two RNA species (5S and 23S rRNA). Although bacteria do not undergo meiosis or mitosis and do not require cellular fusion to initiate reproduction (as bacteria are not diploid), many bacteria do involve a cellto-cell transfer of genomic DNA by various mechanisms. These mechanisms may range from the uptake of exogenous DNA from their environment (a process called transformation) and the integration of a bacteriophage introduces

foreign DNA into the chromosome (a process called transduction), to the acquisition of DNA through direct cell contact (a process called conjugation). The incorporation of genes and DNA from other bacteria or the environment into the recipient cell’s DNA is also called horizontal gene transfer. While DNA transfer occurs less frequently per individual bacterium than that among eukaryotes involving obligate sexual reproduction, the much shorter generation times and high numbers associated with bacteria can make the DNA transfer a significant contributor to the evolution of bacterial populations. Gene transfer is vital to the development of antibiotic resistance in bacteria as it allows the rapid transfer of resistance genes between different pathogens. Regardless of genome size, most organisms show a mutation rate on the order of one mutation per genome per generation. Given their very short generation times (<1 h in culture media and a few hours in the wild) and small genomes (which are a 1000 times smaller than most eukaryotes), prokaryotes generally display 1000 times more mutations per gene, per unit time, and per individual than eukaryotes. Furthermore, with greater population sizes resulting in the absolute amount of mutational variation entering the population, prokaryotes have enormous capacity to adapt to and invade new niches, which are the key factors contributing to the evolutionary success of prokaryotes. Genomic diversity in bacteria comes in two forms: (i) genetic heterogeneity wherein different strains have different alleles of the same gene and (ii) genomic plasticity where different strains have different genes. Recent studies indicate that each strain (serovar) within a bacterial species receives a unique distribution of genes from a population-based supra­ genome that is many times larger than the genome of any given strain.1 Through the autocompetence and autotransformation mechanisms, bacterial strains (or serovars) within the species may evolve and generate diversity in vivo to enable them to persist in the face of myriad host defense mechanisms and environmental stresses. In other words, the strain (serovar)-specific genes (e.g., contingency genes) may provide for an increased number of genetic characters that facilitate the population as a whole to adapt rapidly to environmental factors, such as those experienced in the host during chronic infectious processes. There is evidence that under arduous external conditions, many bacteria form biofilms that often exchange DNA at rates several orders of magnitude greater than planktonic bacteria and that are responsible for many chronic bacterial infections in human patients. For example, biofilm-associated growth of Pseudomonas aeruginosa has been implicated in several chronic suppurative otitis media. It appears that the possession of a distributed genome is a common host interaction strategy.

1.2.5  Ecological and Medical Importance Bacteria play a number of beneficial roles in the ecological balance of our planet. Bacteria are involved in the recycling of carbon dioxide (CO2) to oxygen via photosynthesis;

6

in the decomposition of dead plant and animal matter, improving soil fertility; and in the fixation of nitrogen into the nitrogen compound ammonia for plant growth. Bacteria can help clean up oil spills, pesticides, and other toxic materials. Some bacteria are able to remove (leach) the copper from the ores (copper sulfides), while others are useful for food production such as yogurt, cheese, cider, and vinegar. Some bacteria (e.g., Bacillus thuringiensis or BT, a soil dwelling gram-positive bacterium) can be used as pesticides (trade names Dipel and Thuricide) in the biological control of Lepidopteran pest. In addition, as fast growers with relatively low demands for nutrients, bacteria represent ideal hosts for mass production of certain plastics, enzymes used in laundry detergents, and antibiotics such as streptomycin and tetracycline, as well as pharmaceuticals and fine chemicals upon genetic modification. Furthermore, some bacteria (e.g., Listeria monocytogenes, Mycobacterium bovis Baille Calmette-Guerin or BCG, Salmonella, Shigella, and Escherichia coli) are useful carriers for delivering vaccine molecules against microbial diseases and cancers.2 Although many bacteria are harmless or beneficial, a few can exert detrimental effects on food and plant production, as well as human and animal health. Some bacteria can cause food spoilage (e.g., Lactobacillus) and foodborne diseases (e.g., Shigella, Campylobacter, Salmonella, and Listeria), and others can harm agriculture because of the major diseases of plants and farm animals they cause. For instance, Brachyspira hyodysenteriae causes a type of severe diarrhea in pigs that can have disastrous consequences for pig farmers. Some bacteria are involved in metal corrosion (wearing away) through the formation of rust, especially on metals containing iron. There are approximately ten times as many bacterial cells as human cells in the human body, with large numbers of bacteria on the skin and in the digestive tract. The communities of bacteria and other organisms that inhabit the body are sometimes referred to as the normal microflora or microbiota. Some bacteria in human body produce essential nutrients (e.g., vitamin K) that the body cannot make itself. Nonetheless, some bacteria are highly pathogenic and deadly. Yersinia pestis was responsible for the most fatal and devastating bacterial disease in history—the bubonic plague—which killed an estimated 200 million people prior to the advent of antibiotics. Currently, the most common fatal bacterial diseases are respiratory infections, with tuberculosis (caused by Mycobacterium tuberculosis) alone killing about 2 million people a year. One of the world’s deadliest bacterial diseases today is cholera, which is caused by foodborne Vibrio cholerae. Other globally important bacterial diseases include pneumonia caused by Streptococcus and Pseudomonas, tetanus, typhoid fever, diphtheria, syphilis, and leprosy. Another common bacterial disease is tooth decay, which results from the acids bacteria produce from sugar via fermentation, which dissolves the enamel of the teeth and create cavities (holes) in the teeth.

Molecular Detection of Human Bacterial Pathogens

1.3  BACTERIAL IDENTIFICATION 1.3.1  Current Diagnostic Approaches Identification and characterization of bacteria and their roles in disease processes represent a critical step in the effective management of bacterial infections. Traditionally, bacteria have been identified and diagnosed through examination of their phenotypic characteristics such as Gram staining, cell and colony morphology, and biochemical and serological properties. As a key component in the phenotypic identification scheme, in vitro culture is often time consuming, and its performance is affected by the changing features of bacterial metabolisms. Given the limited variations in bacterial cell and colony morphology, it is impossible to differentiate most bacterial species on morphological criteria alone. Biochemical and serological techniques are helpful in bacterial identification. However, overreliance on these techniques not only adds to the cost of identification, but also creates delay in the result availability. Furthermore, the intrinsic variability of phenotypic procedures remains a potential source of misdiagnosis. In recent decades, DNA- or RNA-based genotypic (molecular) procedures have been increasingly utilized for microbial identification.3 Besides comparison at the genome level (e.g., G + C content determination and DNA–DNA hybridization), detection of nucleotide differences among shared and uniquely present gene regions provides a practical means for bacterial characterization. Due to its crucial roles in cellular function and maintenance, the ribosomal RNA (rRNA) gene is highly conserved (i.e., the least variable) and abundant (with each living cell containing 104 –105 copies of the 5S, 16S, and 23S rRNA molecules). The rRNA gene (and its genomic coding sequence rDNA) therefore offers a valuable target for confirmation of an organism’s taxonomic status and species identity, and for estimation of the rates of species divergence.4–8 In addition, a range of housekeeping-, species-, group-, and virulence-specific genes provide alternative targets for improved determination of bacterial organisms.9,10 The early molecular methods are largely nonamplified, as exemplified by G + C content determination, DNA–DNA hybridization, fluorescence in situ hybridization (FISH), ribotyping, and so forth. Because these methods often require large amounts of starting materials and are cumbersome to perform, they are now rarely used in routine clinical setting, and their applications are limited to the characterization and description of novel bacterial species/subspecies. The more recent molecular methods often involve nucleic acid amplification. These include polymerase chain reaction (PCR), ligase chain reaction (LCR), nucleic acid sequence-based amplification (NASBA), transcription-mediated amplification (TMA), strand displacement amplification (SDA), rolling circle amplification (RCA), cycling probe technology (CPT), branched DNA (bDNA), and loop mediated isothermal amplification (LAMP), and so forth. Due to their efficiency, simplicity, and robustness, PCR and its derivatives have been widely adopted in research and clinical laboratories for

7

Introductory Remarks

specific, sensitive and rapid identification and diagnosis of bacterial pathogens.11 In a standard (conventional) PCR, a pair of oligonucleotides (of about 20 bases in length) is typically used as primers to anneal and amplify a gene region of interest with the help of DNA polymerase. The resulting amplified products (or amplicons) are separated by agarose gel electrophoresis and visualized with a DNA-binding dye (e.g., ethidium bromide). Further refinements of standard PCR led to the development of nested PCR (in which two consecutive PCR are performed one after another), multiplex PCR (in which multiple pairs of primers are included for simultaneous amplification of several genes of interests from the same or different organisms), arbitrarily primed PCR (in which a single oligonucleotide of about 10 bases in length is used to amplify random regions in a genome), and reverse-transcriptase PCR (RT–PCR) (in which RNA instead of DNA is targeted).12,13 Increasingly, the formats of PCR product detection have moved away from gel electrophoresis to enzymatic signal detection (e.g., ELISA and flow cytometry), real-time detection (e.g., using intercalating fluorescent dye [SYBR® green], hydrolysis dual-labeled probes [TaqMan®], hybridization probes [LightCycler], molecular beacons, peptide nucleic acid [PNA] probes, TaqMan minor groove binding [MGB™] probes, locked nucleic acid [LNA®] primers and probes, and scorpions™), line probe assay (LiPA), microarray, sequencing analysis (e.g., pyrosequencing), mass spectrometry (MS), and so forth.14 In particular, real-time PCR demonstrates superior sensitivity, rapidity, and broad dynamic range, eliminates postamplification handling steps (thus minimizing carryover contamination), and is amenable to automation for high throughput detection.15 Furthermore, real-time PCR provides the option for melting curve analysis, permitting discrimination of the amplified product from nonspecific product or primer-dimmers.14 For these reasons, real-time PCR is becoming a method of choice for microbial identification in clinical laboratories worldwide.

1.3.2  Performance Parameters The performance of a diagnostic assay is often evaluated by using several key parameters, including detection limit, sensitivity, specificity, accuracy, intraassay precision, interassay precision, and linearity (as in the case of a quantitative assay). Detection limit (or limit of detection) is the lowest concentration or quantity of bacteria that can be detected by a given assay. Sensitivity is the percentage of samples containing bacteria of interest that are identified by the assay as positive for the bacteria. Specificity is the percentage of samples without bacteria of interest that are identified by the assay as negative for the bacteria. Accuracy (or trueness) is the degree of conformity of an assay’s measurements to the actual (true) value. It is often estimated by analyses of reference materials or comparisons of results with those obtained by a reference method. The closer an assay’s measurements to the accepted value, the more accurate the assay is. Precision is the degree of mutual agreement among a series of assay’s individual

measurements, values, or results. Usually characterized in terms of the standard deviation of the measurements, precision can be stratified into (i) repeatability (the variation arising using the same instrument and operator in a single rune—i.e., intraassay precision—or repeating during a short time period) and (ii) reproducibility (the variation arising using the same measurement process among different instruments and operators from one run to another—i.e., interassay precision—or over longer time periods). Linearity refers to the tendency of measurements by a quantitative assay to form a straight line when plotted on a graph. Data from linearity experiments may be subjected to linear regression analysis with an ideal regression coefficient of 1. In case of a nonlinear curve, other objective, statistically valid methods may be utilized.16

1.3.3  Result Interpretation A positive result by a molecular assay for a given pathogen normally confirms the etiologic relationship if the clinical syndrome is compatible with the pathogen identified. Considering the sensitive nature of the amplified methods such as PCR, it is important to rule out the possibility of a false positive result. Occasionally, false positive results may originate from the low diagnostic specificity of the assay, in which primers bind to irrelevant sequences and occasionally a homologous sequence that is shared among related or unknown bacteria. More often, false positive results in the molecular testing come from contamination, which may arise during manual handling of the samples in the testing laboratory either at the pre or postextraction (while setting up the PCR) stages. This risk is heightened when a high copy number polynucleotide (or plasmid) is used as a quantification standard and distributed around the laboratory, contaminating reaction source. In addition, contamination may be attributable to samples referred from other laboratories that do not utilize manipulation techniques that are mandatory for the molecular testing. These may include the use of unplugged pipette tips, infrequent changing of gloves and using pipette for long periods without decontamination. Another cause of contamination is by amplification products from previous tests.17–19 Contamination may also occur by leakage from tubes or microtiter plates with lids not tightly closed or by breakage of glass capillaries leading to spillage of the amplification mixture. Besides the adoption of stringent laboratory practice, the risk of contamination with PCR products may be reduced by replacing nucleotide dTTP with dUTP in PCR, and implementing a digestion step with UracilDNA-glycosylase (UNG) to remove previous PCR products containing dUTP prior to each amplification reaction. Furthermore, inclusion of multiple negative controls, such as no-template controls (NTC) and no-amplification controls (NAC) may help identify the likely source of contamination and prevent false positive results. Moreover, microbial DNA may come with PCR reagents. Similarly, a negative result by a molecular assay for a given pathogen normally indicates the absence of the pathogen.

8

However, it is equally important to rule out the possibility of false negative results. One possible cause is due to the low sensitivity of the assay employed. Alternatively, an insufficient amount of bacteria may be present in the sample (due to sample degradation or prior antibiotic treatment). Another cause may be the impurity of the processed sample. Enzymes used in PCR and RT–PCR (e.g., DNA polymerase, reverse transcriptase) are impeded by components in blood and feces (e.g., heme, hemoglobulin, lactoferrin, immunoglobulin G, leukocyte DNA, polysaccharides, and urea), in foods (e.g., phenolics, glycogen, calcium ions, fat, and other organic substances), in environmental specimens (e.g., phenolics, humic acids, and heavy metals), and in added anticoagulants (e.g., EDTA and heparin) as well as nucleic acid purification reagents (e.g., detergents, lysozyme, NaOH, alcohol, EDTA, EGTA, phenol, and high salt concentrations).20–24 Any impurities and contaminations present in the samples after nucleic acid isolation may contribute to false negative results. A useful way to determine the effectiveness of a nucleic acid purification procedure for removing inhibitory substances is to spike samples with well-defined DNA or RNA prior to and after sample preparation (as process and amplification internal controls). In light of the high sensitivity of PCR, the occurrence of false negative results is probably a truly underestimated problem. Thus, before definitive diagnoses and treatment decisions are made, all molecular and immunological diagnostic results need be reviewed and critically interpreted in combination with the patient’s clinical presentation.

1.3.4 Standardization, Quality Control, and Assurance Molecular tests such as PCR offer improved sensitivity, specificity, accuracy, precision, and result availability for microbial identification and diagnosis. There is a clear trend toward the adoption and application of these methods in routine diagnostics. However, in view of the possibility of false positive and false negative results occurring in these highly sensitive tests, it is vital that molecular diagnostic methods are properly standardized and validated and appropriate quality control measures are put in place. Standardization and Validation. Standardization of molecular tests addresses the need for standardized reagents and common units, contamination control mechanisms, inhibition control mechanisms, clinically relevant dynamic ranges and internal controls, and so forth. Validation helps to verify the sensitivity, specificity, accuracy, repeatability (intraassay precision), reproducibility (interassay precision), detection limit, and linearity (if quantitative) of molecular tests.25 Before validating a method, it is important to have all instruments calibrated and maintained throughout the testing process. The validation process may involve a series of steps, including (i) testing of dilution series of positive samples (or plasmid construct) to determine the limits of detection of the assay and their linearity over concentrations to be measured in quantitative test (using minimal number of

Molecular Detection of Human Bacterial Pathogens

reference calibrators such as previously tested patient samples or pooled sera); (ii) evaluating the sensitivity and specificity of the assay, along with the extent of cross-reactivity with other genomic material; (iii) establishing the day-to-day variation of the assay’s performance; (iv) assuring the quality of assembled assays using quality control procedures that monitor the performance of reagent batches; and (v) aligning the in-house primer and probe sequences with a genome sequence databank to avoid extended specificity testing.25–28 Quality Control. Quality control strategies for nucleic acid–based tests include (i) designation of a “clean” area for reaction setup (e.g., room under negative air pressure, positive-displacement pipettes, aerosol-block pipette tips, UV-equipped PCR cabinet); (ii) use of personal protective equipment (PPE) (e.g., disposable gloves and lab coats to prevent introduction of contaminating DNA or nucleases); (iii) use of uracil-N-glycosylase (UNG) in real-time PCR (to eliminate crossover amplicon contamination); (iv) use of a “hot-start” method (to minimize false priming events by withholding a crucial reaction component until appropriate temperature is reached); (v) use of external positive and negative controls (to monitor reaction performance and contamination) and homologous or heterologous internal controls (to monitor presence of inhibitors).29 A variety of test controls may be considered for diagnostic PCR. These include (i) internal amplification control (IAC) (negative sample spiked with sufficient pathogen and processed throughout the entire protocol); (ii) processing positive control (PPC) (negative sample spiked with sufficient closely related, but nontarget, strain processed throughout the entire protocol); (iii) reagent control (blank) (containing all reagents, but no nucleic acid apart from the primers); (iv) Premises control (tube containing the master mixture left open in the PCR setup room to detect possible contaminating DNA in the environment [carried out at regular intervals as part of the quality assurance program]); (v) standard (3–4 samples containing a tenfold dilution series of known number of target DNA copies in a range).29–32 Quality Assurance. One way to assess preparedness of the diagnostic laboratories is through the conduct of an external quality assurance (EQA) program that provides characterized specimens containing pathogens of interest. The design of a quality assurance program has the following components: (i) internal quality control (IQC) materials are distributed every month and comprising three pools of clinical samples of known pathogen status (typically one negative, one positive containing 1 log10 over the lower limit of detection of the assay, and one low positive containing up to 1 log10 of the lower limit of detection of the assay). These are incorporated in test runs on a weekly basis. The purpose of IQC is to provide samples of known status for repeated testing in parallel with clinical samples to ensure reproducibility of the test system in an individual laboratory. (ii) EQA distributions of panels of five unknown samples distributed quarterly. Results are returned to the QA laboratory for assessment. EQA compares the performance of different testing sites using specimens of known but undisclosed content. (iii) Aliquots of all samples

9

Introductory Remarks

sent from the reference laboratory are posted back to Site A for repeat testing to check for integrity of the pools and for transport problems. (iv) A final element of the pilot program involves Sites B, C, and D sending an aliquot of every 50th sample to Site A to check for reproducibility. (v) A detailed record of distributions is kept to provide an audit trail.33,34





1.4  CONCLUSION Molecular assays have the potential to increase the speed and accuracy of bacterial identification and diagnosis in research and clinical laboratories. When selecting and implementing a molecular test, a number of factors may be considered. These include (i) cost assessment; (ii) high throughput and automation capability (e.g., 96 or 384 well real-time PCR); (iii) detection limit; (iii) multiple target detection (e.g., multiplex PCR and microarray); (iv) broad range detection (e.g., 16S or 23S rRNA gene sequencing to identify poorly characterized, fastidious or noncultivable bacteria); (v) antimicrobial resistance detection; (vi) ease of use (commercial assays); (vii) sample preparation protocol (commercial kits). To overcome the limitations of molecular tests such as potential false positive and false negative results that impact significantly on patient management, laboratory space must be dedicated for instruments and sample preparation, contamination must be minimized, technicians must have proper training, and quality control procedures must be incorporated into routine laboratory workflow.14 There is a continuing trend toward miniature devices for microbial testing. One such promising device is biosensor. A biosensor incorporates a biological material (e.g., tissue, microorganisms, organelles, cell receptors, enzymes, antibodies, nucleic acids, natural products, etc.), a biologically derived material (e.g., recombinant antibodies, engineered proteins, aptamers, etc.) or a biomimic (e.g., synthetic catalysts, combinatorial ligands and imprinted polymers) in a physicochemical transducer or transducing microsystem (e.g., optical, electrochemical, thermometric, piezoelectric, magnetic, or micromechanical), leading to enhanced detection and identification of microbial pathogens in clinical, food, and environmental specimens.35,36

REFERENCES





1. Hogg, J.S. et al., Characterization and modeling of the Haemophilus influenzae core and supragenomes based on the complete genomic sequences of Rd and 12 clinical nontypeable strains, Genome Biol., 8, R103, 2007. 2. Liu, D., Listeria-based anti-infective vaccine strategies, Rec. Patents Anti-infect. Drug Disc., 1, 281, 2006. 3. Vaneechoutte, M., and Van Eldere, J., The possibilities and limitations of nucleic acid amplification technology in diagnostic microbiology, J. Med. Microbiol., 46, 188, 1997. 4. Jonasson, J., Olofsson, M., and Monstein, H.J., Classification, identification and subtyping of bacteria based on pyrosequencing and signature matching of 16S rDNA fragments, Apmis, 110, 263, 2002. 5. Clarridge, J.E.III, Impact of 16S rRNA gene sequence analysis for identification of bacteria on clinical microbiology and infectious diseases, Clin. Microbiol. Rev., 17, 840, 2004.















6. Barlaan, E.A. et al., Electronic microarray analysis of 16S rDNA amplicons for bacterial detection, J. Biotechnol., 115, 11, 2005. 7. Cherkaoui, A. et al., Development and validation of a modified broad-range 16S rDNA PCR for diagnostic purposes in clinical microbiology, J. Microbiol. Methods, 79, 227, 2009. 8. Sontakke, S. et al., Use of broad range16S rDNA PCR in clinical microbiology, J. Microbiol. Methods, 76, 217, 2009. 9. Liu, D. et al., Characterization of virulent and avirulent Listeria monocytogenes strains by PCR amplification of putative transcriptional regulator and internalin genes, J. Med. Microbiol., 52, 1066, 2003. 10. Liu, D. et al., A multiplex PCR for species- and virulence-specific determination of Listeria monocytogenes, J. Microbiol. Methods, 71, 133, 2007. 11. Lantz, P.G. et al., Biotechnical use of polymerase chain reaction for microbiological analysis of biological samples, Biotechnol. Annu. Rev., 5, 87, 2000. 12. Keer, J.T., and Birch, L., Molecular methods for the assessment of bacterial viability, J. Microbiol. Methods, 53, 175, 2003. 13. Liu, D., Identification, subtyping and virulence determination of Listeria monocytogenes, in important foodborne pathogen, J. Med. Microbiol., 55, 645, 2006. 14. Mothershed, E.A., and Whitney, A.M., Nucleic acid-based methods for the detection of bacterial pathogens: Present and future considerations for the clinical laboratory, Clin. Chim. Acta, 363, 206, 2006. 15. Smith, K., Diggle, M.A., and Clarke, S.C., Automation of a fluorescence-based multiplex PCR for the laboratory confirmation of common bacterial pathogens, J. Med. Microbiol., 53, 115, 2004. 16. Hoorfar, J. et al., Practical considerations in design of internal amplification control for diagnostic PCR assays, J. Clin. Microbiol., 42, 1863, 2004. 17. Millar, B.C., Xu, J., and Moore, J.E., Risk assessment models and contamination management: implications for broad-range ribosomal  DNA PCR as a diagnostic tool in medical bacteriology, J. Clin. Microbiol., 40, 1575, 2002. 18. Aslanzadeh, J., Preventing PCR amplification carryover contamination in a clinical laboratory, Ann. Clin. Lab. Sci., 34, 389, 2004. 19. Borst, A., Box, A.T., and Fluit, A.C., False-positive results and contamination in nucleic acid amplification assays: Suggestions for a prevent and destroy strategy, Eur. J. Clin. Microbiol. Infect. Dis., 23, 289, 2004. 20. Abu Al-Soud, W., and Rådström, P., Capacity of nine thermostable DNA polymerases to mediate DNA amplification in the presence of PCR-inhibiting samples. Appl. Environ. Microbiol., 64, 3748, 1998. 21. Rådström, P. et al., Pre-PCR processing: strategies to generate PCR-compatible samples, Mol. Biotechnol., 26, 133, 2004. 22. Wolffs, P. et al., Impact of DNA polymerases and their buffer systems on quantitative real-time PCR, J. Clin. Microbiol., 42, 408, 2004. 23. Liu, D., Preparation of Listeria monocytogenes specimens for molecular detection and identification, Int. J. Food Microbiol., 122, 229, 2008. 24. Handschur, M. et al., Preanalytic removal of human DNA eliminates false signals in general 16S rDNA PCR monitoring of bacterial pathogens in blood, Comp. Immunol. Microbiol. Infect. Dis., 32, 207, 2009.

10 25. Malorny, B. et al., Standardization of diagnostic PCR for the detection of foodborne pathogens, Int. J. Food Microbiol., 83, 39, 2003. 26. Anon, Microbiology of food and feeding stuffs—protocol for the validation of alternative methods. ISO 16140, first edition, ISO, Geneva, Switzerland, 2003. 27. Josefsen, M.H. et al., Validation of a PCR-based method for detection of foodborne thermotolerant campylobacters in a multi-center collaborative trial. Appl. Environ. Microbiol., 70, 4379, 2004. 28. Hoorfar, J., Wolffs, P., and Radstrom, P., Diagnostic PCR: Validation and sample preparation are two sides of the same coin, Apmis, 112, 808, 2004. 29. Hoorfar, J. et al., Making internal amplification control mandatory for diagnostic PCR, J. Clin. Microbiol., 41, 5835, 2003. 30. Rodriguez-Lazaro, D. et al., Construction strategy for an internal amplification control for real-time diagnostic assays using nucleic acid sequence-based amplification: Development and clinical application, J. Clin. Microbiol., 42, 5832, 2004.

Molecular Detection of Human Bacterial Pathogens 31. Trapmann, S. et al., Development of a novel approach for production of dried genomic DNA standards for quantitative PCR testing of food-borne pathogens, Accred. Qual. Assur., 9, 695, 2004. 32. Burggraf, S., and Olgemoller, B., Simple technique for internal control of real-time amplification assays, Clin. Chem., 50, 819, 2004. 33. Raggi, C.C. et al., External quality assurance program for PCR amplification of genomic DNA: An Italian experience, Clin. Chem., 49, 782, 2003. 34. Cubie, H.A. et al., The development of a quality assurance programme for HPV testing within the UK NHS cervical screening LBC/HPV studies, J. Clin. Virol., 33, 287, 2005. 35. Uslu, F., et al., Label-free fully electronic nucleic acid detection system based on a field-effect transistor device, Biosens. Bioelectron., 19, 1723, 2004. 36. Lazcka, O., Del Campo, F.J., and Munoz, F.X., Pathogen detection: A perspective of traditional methods and biosensors, Biosens. Bioelectron., 22, 1205, 2007.

Section I Actinobacteria

2 Actinomadura Martha E. Trujillo CONTENTS 2.1 Introduction........................................................................................................................................................................ 13 2.1.1 Taxonomy............................................................................................................................................................... 13 2.1.2 Cell Morphology, Physiology, and Isolation........................................................................................................... 14 2.1.3 Differentiation of Pathogen Actinomadura Species............................................................................................... 15 2.1.4 Pathogenicity and Clinical Features....................................................................................................................... 15 2.1.5 Diagnosis................................................................................................................................................................ 17 2.1.5.1 Conventional Techniques......................................................................................................................... 17 2.1.5.2 Molecular Techniques.............................................................................................................................. 17 2.1.6 Treatment of Actinomadura Infections.................................................................................................................. 18 2.2 Methods.............................................................................................................................................................................. 18 2.2.1 Sample Preparation and Observation..................................................................................................................... 18 2.2.1.1 Microscopic Examination........................................................................................................................ 18 2.2.1.2 Isolation and Culture................................................................................................................................ 18 2.2.1.3 Isolation of DNA...................................................................................................................................... 19 2.2.2 Detection Procedures.............................................................................................................................................. 19 2.3 Conclusions and Future Perspectives................................................................................................................................. 20 References.................................................................................................................................................................................... 21

2.1  INTRODUCTION The term actinomycete is an informal designation for filamentous gram-positive bacteria with high G + C content in their genome and that belong to the order Actinomycetales in the class Actinobacteria.1,2 Most aerobic actinomycetes are soil saprophytes and are rarely encountered in clinical practice, but some are serious pathogens of humans and animals causing a number of diseases that include actinomycetoma, actinomycosis, and nocardiosis. The clinical manifestations and severity of the disease and the prognosis in an infected host are extremely variable and may be determined by factors such as the route of infection and the presence or absence of a properly functioning immune system.3 The diagnosis of actinomycete infections has been hindered by a combination of clinical and microbiologic difficulties, including their often nonspecific clinical presentation, a requirement for invasive diagnostic biopsy procedures, difficulty in isolation, and incorrect identification. Strategies to improve outcome for infected patients include a heightened awareness of clinicians and clinical microbiology personnel, which may enable the earliest possible diagnosis; standardization of antimicrobial susceptibility testing methods; and the evaluation of newer effective drug therapies for these patients. In addition, new developments in the

classification of these microorganisms should serve as a framework for the identification of clinically significant species. Actinomycetoma is a localized chronic, destructive, and progressive infection of the skin and subcutaneous tissues caused by aerobic actinomycetes.3–5 The main species involved include Actinomadura madurae, Actinomadura latina, Actinomadura pelletieri, Nocardia brasiliensis, Nocardia otitidiscaviarum, Nocardia transvalensis, Streptomyces somaliensis, and Streptomyces sudanensis.6,7 The disease is endemic in certain tropical and subtropical regions, where it has a devastating effect on patients as it frequently leads to deformities, disabilities, and eventually amputation of the affected organs.7 Actinomycetoma can become dangerous to health, or even life, when treatment is inadequate or delayed. This chapter will focus on the genus Actinomadura and its significance as a pathogen of actinomycetomas and nonmycetomic infections.

2.1.1  Taxonomy The genus Actinomadura contains aerobic, gram-positive, nonacid-fast, nonmotile, and chemo-organotrophic actinomycetes that produce well-developed, nonfragmenting vegetative mycelia and aerial hyphae that differentiate into surfaceornamented spore chains of various lengths (10–50 spores). 13

14

Members of the genus Actinomadura are charac­terized chemotaxonomically by the presence of meso-diaminopimelic acid and madurose in their cell wall with peptidoglycan structures of the acetyl type, major proportions of hexahydrogenated menaquinones with nine isoprene units, complex fatty acid profiles, including hexadecanoic, 14-methylpentadecanoic, and 10-methyloctadecanoic acids as predominant components, and diphosphatidylglycerol and phosphatidylinositol as major phospholipids.8 The systematics of the genus Actinomadura has been significantly improved by the application of modern taxonomic methods.8–14 On the basis of 16S and 23S rRNA gene sequence analyses, the genus is phylogenetically related to members of the family Thermomonosporaceae,1,14 which also includes the genera Actinocorallia, Spirillospora, and Thermomonospora. The family Thermomonosporaceae belongs to the suborder Streptosporangineae in the order Actinomycetales.2 A. madurae was first described in 1894 by Vincent,15 based on several strains isolated from an Algerian case of Madura foot, as “Streptothrix madurae.” The organism was subsequently classified in the genus Nocardia,16 then in the genus Streptomyces.17 The taxonomic status of this microorganism remained controversial until Becker et al.18 found that whole-organism hydrolysates of representative strains contained meso-diaminopimelic acid (meso-DAP) and a characteristic sugar identified as madurose.19 In 1983, compelling evidence that the genus Actinomadura was heterogeneous was provided by Fischer et al.10 when these authors assigned representative strains to two aggregate groups defined on the basis of chemical and nucleic acid pairing data—A. madurae group and Actinomadura pusilla group. The division of the genus Actinomadura into two aggregate groups was formally recognized by Kroppenstedt et al.,8 who proposed that the genus Actinomadura be retained for A. madurae and related species, and that the Actinomadura pusilla group be reclassified in the genus Microtetraspora. Taxonomic work carried out with a group of clinically significant Actinomadura strains received either as A. madurae or A. pelletieri revealed that a third species, A. latina should be officially recognized.12 The genus currently comprises more than 40 validly described species (http://www.bacterio. net),20 but only three—A. latina, A. madurae, and A. pelletieri—are considered pathogens. However, it is becoming increasingly evident that additional pathogenic actinomadurae need to be formally described as new species.6,12,21

Molecular Detection of Human Bacterial Pathogens

of most species form a spore-bearing, powdery aerial mycelium on media such as inorganic salts-starch (ISP 4), oatmeal (ISP 3), and yeast extract-malt extract agars (ISP 2) after cultivation for 10–14 days.22 At maturity, aerial mycelia may be blue, cream, gray, green, pink, yellow, or white, thereby differing little from streptomycetes. Superficial similarity to streptomycetes is also reinforced by the morphology of the sporophores. Actinomadurae and streptomycetes can be distinguished by direct microscopic comparison of cultures. Most Actinomadura strains are conspicuous for the size of their spores. The diameter of spores noticeably exceeds the diameter of the hyphae, whereas in streptomycetes spores and hyphae are of similar diameter. In general, actinomadurae are slow-growing microorganisms. These bacteria generally grow well on modified Bennett’s agar,23 glucose-yeast extract agar,24 and on formulations used for the cultivation of Streptomyces.22 A. latina, A. madurae, and A. pelletieri show moderate-to-good growth on Czapek-dox casamino acids.25,26 Actinomadura strains are strictly aerobic with an oxidative metabolism. Most species grow well between 25°C and 40°C but species such as A. formosensis, A. rubrobrunea, and A. viridilutea are thermophilic and grow between 45°C and 65°C. Actinomadurae can metabolize a wide range of sugars and amino acids as carbon sources for growth. Most strains hydrolyse aesculin, arbutin, casein, elastin, gelatine, testosterone, and Tweens 60 and 80.6,27 Pectinase activity has been reported for A. mexicana, A. napariensis, and A. verrucosispora.6,28 Pathogenic strains of A. madurae produce collagenolytic enzymes.29 Most actinomadurae grow in the presence of NaCl 4% (w/v), but A. atramentaria can grow in NaCl concentrations of 7% (w/v). Diverse culture media have been employed for the isolation of Actinomadura strains, especially from soil samples. The

2.1.2  Cell Morphology, Physiology, and Isolation Actinomadura strains characteristically form nonfragmenting, extensively branched, substrate mycelia and aerial hyphae, which carry from 1 to 50 arthrospores. Spores are borne in curled, hooked, spiral, or straight chains and the spore surface may be folded, irregular, rugose, smooth, shiny, or warty. Some strains, notably those from clinical sources, mostly lack aerial mycelium, with colonies exhibiting a cartilaginous or leathery appearance (Figure 2.1). However, members

FIGURE 2.1  Actinomadura sp. grown on ISP 2 agar for 10 days. The strain does not produce aerial mycelium or diffusible pigments.

Actinomadura

media that have proved most suitable include oatmeal agar (ISP 3), yeast extract-malt extract agar (ISP 2), starch-mineral salts agar (ISP 4), Bennett-sucrose agar, and glycerol-asparagine agar (ISP 5). Strains may be isolated from agar plates by dilution techniques after incubation for up to 6 weeks. Trujillo and Goodfellow6 successfully isolated new Actinomadura species from environmental samples collected in Hong Kong, Kenya, Mexico, South Africa, and Venezuela using the selective isolation procedure recommended by Athalye et al.24 Pathogen Actinomadura species can be isolated from clinical samples, including pus and biopsy material, using brain heart infusion agar,30 Sabouraud dextrose agar,31,32 and Columbia agar supplemented with 5% sheep blood.33 All samples should be incubated aerobically between 27°C and 37°C for up to 3 weeks and examined both macroscopically and microscopically for growth every 2 days. Actinomadurae can be recognized by their filamentous appearance, leathery colonies, and by the production of red prodiginine pigments. A. madurae and A. pelletieri produce prodigiosin-like pigments34,35 that are similar to those of Serratia marcescens. Members of these species isolated from patients produce prodiginines characterized by a tripyrrole skeleton and identified as cyclononylprodiginine, nonylprodiginine, and undecylprodiginine. A. latina, A. madurae, and A. pelletieri strains are mainly isolated from clinical material, though there is some evidence that members of A. madurae are widespread in organically rich soils. A. madurae strains isolated from environmental samples tend to lack the red endopigment of clinical isolates and sporulate more readily.34–36

2.1.3  Differentiation of Pathogen Actinomadura Species Published descriptions of Actinomadura species are often incomplete since different investigators emphasize some phenotypic features and omit others, thereby making identification difficult. Nevertheless, most species can be separated using a combination of morphological and physiological properties, though in most cases only the type strain has been studied. However, even when several strains have been studied (e.g., A. madurae, A. pelletieri), the results tend to be variable or inconsistent when those from the literature are compared. Enzymatic substrates based on the fluorophores 4-methylumbelliferone (4-MU) and 7-amino-methylcoumarin (7-AMC) were carried out by Trujillo and Goodfellow6,12,36 to differentiate Actinomadura species. Encouraging results were obtained for differentiating between pathogenic Actinomadura species (Table 2.1).6,36 Enzymatic tests were highlighted by Wink et al.37 to differentiate A. kijaniata from A. namibiensis, as they share a DNA homology higher than 70%.

2.1.4  Pathogenicity and Clinical Features Actinomycetoma. Actinomycetoma is a localized chronic, destructive infection of the skin and subcutaneous tissues

15

caused by aerobic actinomycetes, Actinomadura being one of the main causal agents. The disease is characterized by  progressive swelling of the infected area, distortion of the normal anatomy, and multiple draining sinuses and fistulae.3–5,38–40 Purulent discharge containing the causative agent in the form of grains is characteristic of advanced stages of the disease. The grains vary in size and consistency and may be white, yellow, brown, red, or black depending on the causative agents. Almost 80% of the infections are through the lower extremities of the body; this probably explains the etiology of “Madura foot” described by Gill of Madura in South India in 1842. In areas where actinomycetoma is endemic, it is a common habit to walk barefooted, exposing the skin to thorns and splinters on the soil; as a result, natural infection appears to be more frequent. Other sites of the human body may also be affected such as the back, head, knee, arm, and neck.41,42 In Mexico, the back is the second most common location.43 The clinical picture of actinomycete mycetoma is almost uniform irrespective of the causal agent. In contrast to eumycetoma (caused by eumycetes), infections caused by actinomycetes are more aggressive and destructive and involve both muscle and bone at an early stage.5,41 In general, actinomycetoma is painless, even when the disease is at an advanced stage.41 However, respiratory, neurologic, or other symptoms may be present when the disease affects the chest, head, and neck, or spine. The invasion of the periosteum and adjacent bones may lead to osteomyelitis.42 The infection remains localized and constitutional disturbances are rare, but when they do occur they are generally due to septicemia or to immune depression. Actinomycetoma can produce many disabilities, distortion, and deformity. It can be fatal especially if it affects the skull.41,44 Nonmycetomic Infections. Nonmycetomic infections produced by A. madurae have been reported, including one involving an immunocompromised patient.45 These observations suggested that A. madurae could play a role in pneumonitis or bronchitis. Although no detailed epidemiological studies have been carried out, Bar et al.46 reported a growing number of patients with pneumonia caused by A. madurae. Most of the patients had been infected as a consequence of impaired immunity. These studies suggested that A. madurae should also be considered as a causative agent for nonmycetomic infections such as pneumonia. Since that time, no epidemiological studies on Actinomadura strains from nonmycetomic regions or clinical specimens such as sputum, bronchoalveolar lavage, and blood samples have been reported. However, in Japan between 1996 and 2004, 21 actinomycete strains were isolated from sputum and broncho alveolar lavage.21 Molecular identification using 16S rRNA gene sequencing indicated that all strains belonged to the genus Actinomadura but were unrelated to A. madurae or A. pelletieri. Ninety-five percent of the clinical isolates showed a close phylogenetic relationship with the species Actinomadura cremea and Actinomadura nitritigenes, while the remaining strains could not be related to any other species due to their moderate 16S rRNA gene sequence similarity.

16

Molecular Detection of Human Bacterial Pathogens

TABLE 2.1 Differential Characteristics That Separate Pathogenic Actinomadura Species A. latina

Species

A. madurae

A. pelletieri

Enzyme tests Cleavage of 7-amino-4-methylcoumarin (7AMC-) substrates

Glutaryl-l-phenylalanine-7AMC l-pyroglutamide-7AMC

−

+

+

+

+

−

−

d

+

+

+

−

+

+

−

–

+

−

+

+

−

+

+

−

–

+

−

−

+

−

+

+

−

−

+

−

+

+

−

+

+

−

+

+

−

+

+

−

+

−

−

+

+

−

+

+

−

+

−

+

Cleavage of 4-methylumberlliferone (4MU-) substrates

4MU-N-acetyl-d-galactosoamide 4MU-α-l-arabinopyranoside 4MU-β-d-cellopyranoside 4MU-β −d-galactoside 4MU-β-d-glucoside 4MU-β-d-glucuronide 4MU-β-d-lactoside 4MU-sulfate 4MU-xyloside Growth on sole carbon sources (1%, w/v)

Adonitol Fructose Galactose Glycerol Mannitol Melezitose Degradation tests

Arbutin Esculin Starch

Source: Based on data from Trujillo, M.E., and Goodfellow, M., Antonie van Leeuwenhoek, 84, 39, 2003 and Trujillo, M.E., and Goodfellow, M., Zbl. Bakt., 285, 212, 1997.

Epidemiology. It is not possible to state with any certainty the prevalence of Actinomadura infections (mainly actinomycetoma) in any part of the world, including the endemic areas, since there have not been any international surveillance efforts to ascertain the incidence of the disease. At most, information is limited to a few studies that have been made by individual researchers.21,40,41,47 Actinomycetomas have a worldwide distribution but occur mainly in the tropical and subtropical regions in the area between the latitudes of 15°S and 30°N. This area includes countries such India, Mali, Mexico, Senegal, Somalia, Sudan, and much of Central and South America.5 Sudan appears to have the highest number of actinomycetomas in the world followed by Mexico with an average of 70 cases per year.40 In these countries, the actinomycetoma infections are a major health problem, commonly affecting farmers and herdsmen. In general, the disease is four times more frequent in males than females and mainly affects adults between the ages of 16 and 40 years. Since these are the most active members of the society, the disease has a socioeconomic effect on dependent family members in underdeveloped countries.48 Nevertheless,

cases involving children and elderly men do occur.49–51 Actinomycetoma infections have also been reported in the United States and Europe although in most cases, the patient contracted the disease in a different country.38,52 It is interesting to note that actinomycetoma becomes more active and aggressive in pregnant women. Hormonal changes and decreased immune response during pregnancy may be the explanation for this observation.41 There is evidence that climatic conditions may play a major role in the distribution of actinomycetomas. In the case of Mexico and Sudan, these countries have a rainy season from June to October; a dry, cool season from October to March; and hot and dry weather without rainfall from March to June. In Venezuela, most cases of actinomycetoma have been reported from semiarid areas colonized by trees, bushes, and cacti with thorns. The microorganisms are usually present in the soil. Traumatic inoculation of the subcutaneous tissues caused by sharp objects such as thorns or splinters is thought to be the route of entry. However, this theory has been recently disputed, as many patients have no history of trauma at the infection site.41

Actinomadura

2.1.5  Diagnosis The clinical diagnosis of actinomycetoma becomes more apparent in advanced disease with the development of characteristic sinuses and discharging grains; thus, the disease is often at an advanced stage when diagnosed. 2.1.5.1  Conventional Techniques Microscopic Observation and Isolation. The diagnosis of the actinomycetoma is currently based on the isolation and identification of the causal microorganisms. In practice, the isolation of the causal agent is not always possible or can take a long time. Direct microscopic examination of the pus from the lesions in the actinomycetomas with 10% KOH or saline reveals the presence of granules. The size, form, and color, together with the presence or absence of clubs or pseudoclubs, gives a clue to the identity of the etiologic agent. In the case of A. madurae, the granules can be seen without the aid of a microscope; in other species, the granules are smaller.42 Isolation of the microorganisms can be achieved by culture of the pus, granules, or tissue samples using various culture media such as Sabouraud, mycobiotic, or blood agar media.33,42 The culture technique is often cumbersome and time consuming, and possible sample contamination may give a false positive result. This technique also requires experience to identify the causative microorganisms. Fine Needle Aspiration Cytology. Actinomycetoma can be accurately diagnosed by fine needle aspiration cytology.53–55 Mycetoma lesions have a distinct appearance in a cytology smear and are characterized by the presence of polymorphous inflammatory cells such as neutrophils, histiocytes, lymphocytes, plasma cells, macrophages, and foreign body giant cells. This allows differentiation from artifacts and inflammatory lesions caused by other bacteria and fungi. Diagnosis of actinomycetoma using fine needle aspiration cytology has proved to be as accurate as histopathological observations when the grains are present.54 This technique is fast, inexpensive, and easy to apply, and it is well tolerated by patients. It can be used in routine diagnosis and as an effective means of collection of material for culture and immunological studies. Due to the simplicity of the technique, it can be used in epidemiological surveys of mycetoma and for detection of early cases when radiological and serological techniques may not be useful.41 Radiology. The radiological features of mycetoma are complex. As the disease spreads to the bone, the earliest changes include periosteal erosion and adjacent sclerosis. There may be a soft tissue mass with obliteration of fascial planes. Bone cortex may be compressed from outside by the mass, producing scalloping, which is followed by variable amount of periosteal reaction. Sunray appearance and Codman’s triangle may be present, producing a picture similar to that of osteogenic sarcoma. There may be multiple cavities within normal-density bone; these cavities are small and numerous and their edges not well defined. The cavity size is directly related to the size of the grains. Osteosporosis

17

is common in advanced mycetoma due to compression of the bone and insufficient blood irrigation caused by pressure on the blood vessels. Radiographs are helpful to determine the extent of the infection, although they are not diagnostic. Computed tomography or magnetic resonance imaging can also be helpful to determine the full extent of bone involvement and to delineate soft tissue involvement.39 Immunodiagnosis. Enzyme-linked immunosorbent assays have been used to screen for antibodies against S.  somaliensis and A. madurae in two regions of Sudan.56 These results indicated that a relatively large proportion of clinically asymptomatic individuals in one region were infected with S. somaliensis and, to a lesser degree, A. madurae. To date, however, there are no specific and reliable serologic or immunologic tests useful in the diagnosis of actinomycetoma. This is in part because of the lack of specific antigens that do not cross-react with antibodies of infections caused by other related microorganisms.42 Attempts have been made to develop a delayed-type hypersensitivity test in mycetoma using microbial agents, but in most cases the assays have not been sensitive enough, or represented, as in the case of aerobic actinomycetic infection, cross-reactions with tuberculosis, and leprosy.42 2.1.5.2  Molecular Techniques PCR-RFLP Analysis. Use of PCR coupled with restriction endonuclease and/or probe hybridization analyses of PCR products has been the focus of recent interest for the separation of mycobacteria from nocardiae as well as for the recognition of species within the genera Mycobacterium and Nocardia.57 The method has proven to be sensitive, less time consuming, and less labor intensive than traditional biochemical methods for the identification of the clinically significant species. This technique can be directly applied to clinical isolates provided that sufficient biomass is available for genomic DNA extraction. A 439-bp fragment corresponding to the 65-kDa heat shock protein gene is then amplified by PCR using primers TB11 and TB12,57 and the PCR product is then digested using a combination of five commercially available restriction endonucleases (BstEII, HaeIII, MspI, HinfI, and BsaHI). After digestion, the restriction fragments are electrophoresed on 3% Metaphor agarose containing ethidium bromide at 95 V for 1.5–2 h to obtain the RFLP band patterns. This PCRRFLP methodology distinguished clinical isolates of aerobic actinomycetes, including A. madurae, with 96.8% accuracy. Thus, identification of clinical isolates can be accomplished within 24–48 h of receipt of pure cultures and therefore this system can be readily and economically implemented for routine clinical use.57 16S rRNA Gene Sequencing. Identification of Actino­ madura actinomycetes from nonmycetomic clinical samples has also been carried out successfully using 16S rRNA gene sequencing. In this case, Hafany et al.21 isolated Actinomadura strains from the sputa or the bronchoalveolar lavage fluid of patients with pulmonary infections.

18

For DNA sequencing, bacterial strains can be cultured in brain heart infusion (BHI) broth for 4 days at 32°C. Preparation of genomic DNA samples for PCR amplification can be achieved by applying almost any protocol described in the literature for bacterial DNA isolation. In some cases, crude cell extracts are sufficient to obtain DNA for PCR amplification. Thus, a cell suspension can be heated in boiling water for 10 min and a sample of the supernatant is used directly for PCR.58 16S rRNA genes can be amplified by PCR and directly sequenced using universal primers widely available in the literature.14,58 The obtained sequences can then be compared to those deposited in public databases (e.g., EMBL, NCBI, Genbank, etc.) for identification at the genus level of the microorganism. A useful tool is the EzTaxon.org server,59 which contains a manually curated 16S rRNA database of type strains of prokaryotes, providing a workframe where nontype strains and environmental clones, which in many cases contain erroneous and mislabeled sequences, have been excluded. Therefore, sequence comparison and identification will only be performed with validated sequences.

2.1.6  Treatment of Actinomadura Infections Medical Treatment. The most beneficial treatment begins at an early stage and consists of surgical debridement in conjuction with prolonged antibiotic medications. Combination drug therapy appears to achieve the best results because of a synergistic effect and reduces the likelihood of resistance.39 Treatment periods are characteristically long, especially with bone and visceral involvement, and depend on clinical response. Cure may be defined by a lack of clinical activity response. Several drugs have been used in the treatment of actinomycetoma caused by Actinomadura species, including streptomycin, dapsone, amikacin, and trimethoprim-sulfamelthoxazole.60 Boiron et al.61 used the disk diffusion method to assess the susceptibility of the pathogen species A. madurae and A. pelletieri against 29 different antibiotics. Favorable in vitro susceptibilities were obtained particularly with amikacin, imipenem, aminocycline, tobramycin, and vancomycin. The addition of aminoglycosides to cotrimoxazole gives better efficacy, and is associated with shorter treatment duration. An extensive study on antibiotic susceptibility of the genus Actinomadura was carried out by Trujillo and Goodfellow.6 Many strains were resistant to ampicillin and rifampicin. On the other hand, with the exception of A. oligospora and A. yumaensis, most strains studied were susceptible to tetracycline. Vera-Cabrera et al.62 examined 24 A. madurae strains isolated from patients with actinomycetoma for their sensitivity to new quinolones, including garenoxacin, gatifloxacin, moxifloxacin, and two oxazolidinones: linezolid and the compound DA-7867, and found that all but one of the strains showed high susceptibility to all of these agents; they were

Molecular Detection of Human Bacterial Pathogens

most active to DA-7867, showing a minimum inhibitory concentration of 0.06 µg/mL. However, further work needs to be done to determine whether these results can be extrapolated to antibiotic activity in subcutaneous abscesses and granulomas that are formed in cases of mycetoma. Surgical Treatment. Surgical excision of early and welldefined lesions of mycetomas carries the best prognosis. Drug treatment is given before surgery to decrease lesion size, and continued afterwards to ensure complete clearance of infection. Nonetheless, relapse rates of 20%–90% have been reported. Where there is bone involvement, amputation of the affected limb often becomes unavoidable but must be an absolutely last resort. Unfortunately, in resource-poor regions where there is no available effective drug treatment, and where bone involvement may be complicated by secondary bacterial infection, it is sometimes the only option available to both doctors and patients.41,42

2.2  METHODS 2.2.1  Sample Preparation and Observation Noncontaminated clinical samples including bone tissue, bronchial washings, cerebrospinal fluid, sinus discharge, urine, biopsy, and autopsy material can be collected for examination of the presence and isolation of Actinomadura. Clinical material, especially of sputa and sinus discharge, should be examined as soon as possible to prevent overgrowth by contaminants. If storage is unavoidable, samples should be kept at 4°C without the addition of preservative. Fluid material (e.g., cerebrospinal, urine, blood, etc.) can be examined in wet mounts under the microscope without staining. Samples should be centrifuged (× 4000 rpm) and examined between slide and coverslip for the presence of branching filaments using greatly reduced light or phase contrast. Material from actinomycetomas usually contains granules that facilitate an immediate presumptive identification of the causative agent and provide a good source for isolation of the microorganism. 2.2.1.1  Microscopic Examination



1. Prepare a Gram stain smear from fresh material, from sediment after centrifugation, or from granules crushed between two slides. Gram-positive branched filaments (ca. 1 µm in diameter) are seen at high magnification (×1000). 2. Stain a second smear and examine for acid fastness. Actinomadurae are nonacid-fast.

2.2.1.2  Isolation and Culture Clinical samples should be inoculated in media such as brain heart infusion (Oxoid CM261), Sabouraud dextrose,31 and yeast extract agars24 (see also section on isolation). Media should be transparent to facilitate direct microscopic examination of colonies. The grains should be cleaned several times in sterile saline and crushed with a sterile glass rod to obtain material for inoculation. Several plates should be

Actinomadura

streaked and incubated at 30°C–37°C. Colonies may grow after 7–10 days, but longer incubation periods are often necessary. Actinomadurae can be recognized by their filamentous appearance, leathery colonies, and by the production of red prodigiosin pigments. The colonies are usually folded, with an irregular surface and present a hard texture. Colony color may vary depending on the species, A. madurae and A. latina colonies are usually yellow-white or pink, while A. pelletieri colonies are red or purplish. 2.2.1.3  Isolation of DNA There are many DNA extraction commercial kits available to isolate DNA for PCR amplification, however, DNA crude extract preparations also work well. We present two DNA extraction protocols that have been successfully applied to obtain DNA templates from Actinomadura strains. Method 1. Strains of Actinomadura can be grown on Mueller-Hinton agar or in Mueller-Hinton broth supplemented with 0.4% Tween 80% and 10% oleic acid albumin-dextrose solution. Alternatively, the strains can be grown on any of the media mentioned in Section 2.1.2. The cells are harvested and suspended in 10 mM Tris-1 mM EDTA (pH  8.0) to a density equivalent to that of a MacFarland no. 4 standard, inactivated by submersion in a water bath at 80°C for 30 min, and disrupted for 5 min with zirconium beads (0.1–0.15 mm in diameter) in a Mini-BeadBeater. The resulting ground cell supernatants are subsequently clarified by centrifugation (7500 × g for 5 min). The supernatant is transferred to a clean tube and stored at 4°C.63 ‌ Method 2. Actinomycete strains are grown in 10 mL of tryptone yeast extract broth (ISP 1)22 with agitation at 120 rpm for 24–48 h. Five mL of cultured cells are harvested by centrifugation (7500 × g for 5 min), washed once with 500 µL of TE buffer (10 mM Tris-HCl 1 mM EDTA pH 7.5), and resuspended in 500 µL TE buffer. The samples are heated in boiling water for 10 min, allowed to cool for 5 min and centrifuged (7500 × g for 5 min). The supernatant (300 µL) is transferred to a clean microtube and store at 4°C. If melanin or pigments are produced in ISP 1 broth, the microorganisms can be grown in Middlebrook 7H9 broth, as pigments may interfere with the PCR reaction.

2.2.2  Detection Procedures (i) PCR Detection  The application of PCR-based protocols has become and indispensable tool to aid in the identification of bacteria. Specifically, 16S rRNA gene sequencing provides and accurate and rapid way to identify clinically significant aerobic actinomycetes including Actinomadura. Procedure  A 50 µL reaction mixture can be prepared as follows: 5 µL PCR reaction buffer (10×, PerkinElmer); 3 µL MgCl2 (25 mM); 2 µL bovine seroalbumin (0.1% p/v, Sigma); 0.5 µL primer SF1 (20  µM) (5′-AGAGTTTGATCMTG GCTCAG-3′); 0.5 µL primer SR5 (20 µM) (5′-AAGGAGGT GWTCCARCC-3′); 1.25 µL dNTP mix (10 mM each); 0.8 µL Gold Taq polymerase (5 U/µL, PerkinElmer); DNA

19

template (4 µL), sterile MilliQ water (33 µL). The PCR amplification is performed with an initial denaturation step at 95°C for 9 min, followed by 35 cycles with denaturation at 94°C for 1 min, annealing at 55°C for 1 min, and extension at 72°C for 2 min, with a final extension of 72°C for 7 min. PCR products are then electrophoresed on 1% agarose gels containing ethidium bromide (10 mg/mL) to ensure that the correct fragment has been amplified (~1500 bp). PCR fragments are subsequently excised from the agarose gel and purified using QIAquick PCR purification kit (Qiagen). Sequence reactions are performed on an ABI377 sequencer (Applied Biosystems) using a BigDye terminator v3.0 cycle sequencing kit as supplied by the manufacturer. The following primers are used to obtain the complete 16S rRNA gene sequence (corresponding positions in the E. coli 16S rRNA gene sequence are given in parenthesis): (342); SR2 5′-GWATTACCGCGGCKGCTG-3′ (519); SR3, 5′-CCGTCAATTCMTTTRAGTTT-3′ (907); SR4, 5′-GGGTTGCGCTCGTTG-3′ (1100) and SR5, 5′-AAGGAGGTGWTCCARCC-3′ (1525). The assembled sequence is introduced in the public databases (e.g., GenBank/EMBL) for identification. Given the fact that GenBank/EMBL databases also contain a huge amount of environmental and uncultured bacterial sequences, it is sometimes difficult to obtain a clear identification of the microorganism if the user is not very experienced using these databases. To overcome this difficulty, we highly recommend to use the Eztaxon server (http://www.eztaxon. org), which offers a fully updated database of type strains with validly published prokaryotic names.59 Note 1. In many cases, identification at the genus level can be obtained by sequencing a hypervariable region within domain 1 of the gene using primer SR2. The ~450 bp sequenced DNA fragment is in most cases sufficient for correct identification of strains at the genus level. Note 2. Couble et al.64 have developed a 16S PCR-based assay for the rapid detection of Nocardia spp. directly from human clinical specimens such as skin biopsy samples, pus from abscesses, sputa, or bronchalveolar liquid. Although this method has not been used for the detection of Actinomadura, a similar approach could be applied if genus specific primers for Actinomadura are designed. Below we describe the published method for Nocardia. Sample preparation for direct detection is prepared as follows: Resuspend 100 µL pus samples or 100 mg tissue biopsy specimens in 250 µL of sterile water (molecular grade). This mixture is incubated for 3 h at 55°C with proteinase K at 20  mg/mL (Sigma Aldrich) and inactivated for 15 min at 95°C. Nocardial DNA is then extracted with an MTB respiratory specimen preparation kit (Roche), according to the manufacturer’s instructions. PCR amplification of a 16S rRNA gene fragment using Actinomadura specific primers could then be performed. PCR conditions given above can be used to amplify these fragments taking into consideration the Tm of the primers designed to optimize the annealing temperature.

20

(ii) Genotyping  Protocol 1. PCR-restriction fragment length polymorphism (PCR-RFLP) of the 65-kDa heat shock protein gene. PCR Amplification. Method 1 is recommended for DNA extraction. A 439-bp segment of the 65-kDa heat shock protein (HSP) is amplified from ground cell supernatants by PCR with 1.0 U of Taq DNA polymerase (Boehringer) in optimized buffer E (1.5 mM MgCl2 [pH 9.0]; Invitrogen) containing 42 µM (each) dNTP mixture, 9% dimethyl sulfoxide, and 1 µM (each) primers TB11 (5′-ACCAACGATGGTGTGTCCAT-3′) and TB12 (5′-CTTGTCGAACCGCATACCCT-3′). PCR conditions are 45 cycles at 94°C, 55°C, and 72°C for 1 min each and then for a 10 min final extension at 72°C. Positive and negative controls should be included for comparison. RFLP Analysis. Enzymatic digestion using commercially available restriction endonucleases MspI and HinfI is performed at incubation temperatures and using the buffers at the 10× concentration as recommended by the manufacturers. Restriction fragments are subsequently electrophoresed on 3% MetaPhor agarose containing ethidium bromide (0.625 µg/mL) in a Mini-Sub Cell electrophoresis system at 95V for 1.5–2.0 h. The PCR product digested with MspI DNA yields a two-band profile (265/180 bp). The PCR product digested with HinfI does not present any recognition sites. This combination should be sufficient to identify Actinomadura strains. However, it is very important to compare with appropriate controls. Note. A timed protocol of PCR-RFLP has been presented by Wilson et al.65 where by a clinical isolate can be identified within 2–3 working days. Protocol 2. Amplified rRNA restriction analysis (ARDRA) PCR Amplification. Method 2 is recommended for DNA extraction. PCR is carried out in 50 µL volumes containing 2 mM MgCl2, 2 U Taq ­polymerase (Roche), 150 µM (each) dNTPs, 0.5 µM of each primer, and 2 µL template DNA. Pri­ mer F1 is 5′-AGAGTTTGATCITGGCTCAG-3′, I = inosine and primer R5 is 5′-ACGGITACCTTGTTACGACTT-3′. The primers amplify the nearly full-length 16S rRNA gene. PCR conditions: initial 96°C for 2 min; 30 cycles of 96°C for 45 s, 56°C for 30 s and 72°C for 2 min; and a final 72°C for 5 min. The PCR products are then electrophoresed on 1% agarose gels, containing ethidium bromide (10 g/mL), to ensure that a fragment of the correct size is amplified. PCR products are purified using the QIAquick PCR ­purification kit (Qiagen). Resctriction Endonuclease Digestions and Analysis. The purified 16S rRNA gene should be digested using three different endonucleases, Sau3AI, AsnI, and PstI. Digestion reactions are incubated at 37°C for 3–4 h. Samples are then electrophoresed on 1.5% agarose gels containing ethidium bromide. DNA digested with Sau3AI yields a large fragment of DNA between 980 and 1300 bp. The 16S rRNA gene should not be cut by AsnI. Digestion of the 16S rRNA gene with PstI yields a band of 400–465

Molecular Detection of Human Bacterial Pathogens

bp. The combination of the three results should identify the strain to the genus Actinomadura.58

2.3  C  ONCLUSIONS AND FUTURE PERSPECTIVES Considerable challenges remain in the management of actinomycete infections, especially actinomycetomas. Unlike well-known tropical diseases such as leishmaniasis, malaria, and leprosy, actinomycetomas have received comparatively little international recognition. As a result, there has been little impetus for a new drug development, although they represent a group of tropical infections for which existing treatment options are characterized by high levels of inefficacy. Correct species identification is important for initial diagnosis, treatment evaluation, and epidemiological studies. Modern molecular techniques such as PCR technology are valuable tools for identifying causative species where culture methods have failed. The availability of good drug-susceptibility testing is important given the high rates of resistance to commonly used first-line agents. Actinomycetomas are managed by a combination of antibiotic treatment, mainly sulphonamides, aminoglycosides, or tetracyclines. Carbapenems and oxazolidinones are promising treatment options. Nonetheless, in countries with significant disease burden, such as Mexico, Sudan, and India, physicians have used novel drug combinations in efforts to overcome drug resistance and improve efficacy. However, studies have not been randomized, species identification has not always been possible, clinical endpoints are not accurately stated, and follow-up periods are not defined.43 Furthermore, the vast majority of studies have small patient numbers. Larger, multicenter trials are needed in order to better evaluate drug efficacy and safety. There are also relatively few published studies of current treatment regimes, and therefore they too need to be optimized by randomized blinded trials. New drugs will need to be made affordable to local populations, as this is a disease that primarily afflicts those of low socioeconomic status. Efforts continue to improve molecular diagnostic techniques that are not only sensitive and specific but also might readily be deployed in resource-poor settings. The availability of good drug-susceptibility testing and development of prognostic markers will also improve disease management. Insufficient information is available on the pathogenicity mechanisms of Actinomadura, mainly due to lack of studies. Much work is also needed in this area. Finally, Actinomadura infections, especially actinomycetoma, can be treated given the presence of properly trained physicians, medical students, nurses, and paramedical staff in countries where these infections are endemic. It is also essential to provide health education to farm workers so that they can be protected. It is, however, important that rapid and efficient techniques are developed to identify the causal agents and to provide correct therapeutic treatment.

Actinomadura

21

REFERENCES

19. Lechevalier, M.P., and Gerber, N.N., The identity of madurose with 3-O-methyl-galactose, Carbohydr. Res., 13, 451, 1970. 20. Euzéby, J.P., List of bacterial names with standing in nomenclature: A folder available on the Internet, Int. J. Syst. Bacteriol., 47, 590, 1997, (List of Prokaryotic names with standing in nomenclature. http://www.bacterio.net). 21. Hafany, A. et al., Majority of Actinomadura clinical isolates from sputa or bronchoalveolar lavage fluid in Japan belongs to the cluster of Actinomadura cremea and Actinomadura nitritigenes, and the description of Actinomadura chibensis sp. nov, Mycopathologia, 162, 281, 2006. 22. Shirling, E.B., and Gottlieb, D., Methods for characterization of Streptomyces species, Int. J. Syst. Bacteriol., 16, 313, 1966. 23. Jones, K.L., Fresh isolates of actinomycetes in which the presence of sporogenous aerial mycelia is a fluctuating characteristic, J. Bacteriol., 57, 141, 1949. 24. Athalye, M., Lacey, J., and Goodfellow, M., Selective isolation and enumeration of actinomycetes using rifampicin, J. Appl. Bacteriol., 51, 289, 1981. 25. Cross, T., and Attwell, R.W., Recovery of viable thermoactinomycete endospores from deep mud cores. In Barker, Gould and Wolf (Editors), Spore Research 1973, p. 11, Academic Press, London, 1974. 26. Trujillo, M.E., Taxonomic revision of the genus Actinomadura and related taxa using rapid methods, Ph.D. thesis, University of Newcastle, Newcastle upon Tyne, England, UK, 1994. 27. Kroppenstedt, R.M., and Goodfellow, M., The family Thermomonosporaceae. In Starr, Stolp, Trüper, Balows, and Schlegel (Editors), The Prokaryotes: A Handbook of Habitats, Isolation and Identification of Bacteria, p. 1085, SpringerVerlag, Berlin, 1992. 28. Cook, A.E., le Roes, M., and Meyers, P.R., Actinomadura napierensis sp. nov., isolated from soil in South Africa, Int. J. Syst. Evol. Microbiol., 55, 703, 2005. 29. Rippon, J.W., Extracellular collagenase produced by Streptomyces madurae, Biochim. Biophys. Acta, 159, 147, 1968. 30. Schaal, K.P., Zur mikrobiologisher Diagnostik der Nocardiose, Zentralbl. Bakteriol. Parasitenkd. Infektionskr. Hyg. I. Abt. Orig., 220, 242, 1972. 31. Gordon, M.A., Aerobic pathogenic Actinomycetaceae. In Lennette, Spauling, and Truant (Editors), Manual of Clinical Microbiology, 4th Ed., p. 175, American Society for Microbiology, Washington D.C., 1974. 32. Maiti, P.K. et al., Mycetoma caused by a new red grain mycetoma agent in two members of a family, J. Postgrad. Med., 49, 322, 2003. 33. De Palma, L. et al., A rare european case of madura foot due to actinomycetes, Joint Bone Spine, 73, 321, 2006. 34. Gerber, N.N., Prodigiosin-like pigments from Actinomadura (Nocardia) pelletieri, J. Antibiot., 24, 636, 1971. 35. Gerber, N.N., Minor prodiginine pigments from Actinomadura madurae and Actinomadura pelletieri, J. Heterocycl. Chem., 10, 925, 1973. 36. Goodfellow, M., Trujillo, M.E., and Alderson, G., Approaches towards the identification of sporoactinomycetes that cause mycetoma, Biotechnologia, 7–8, 271, 1995. 37. Wink, J. et al., Actinomadura namibiensis sp. nov, Int. J. Syst. Bacteriol., 53, 721, 2003.

















1. Stackebrandt, E., Rainey, F.A., and Ward-Rainey, N.L., Proposal for a new hierarchic classification system, Actinobacteria classis nov. Int. J. Syst. Bacteriol., 47, 379, 1997. 2. Zhi, X.-Y., Li, W.-J., and Stackebrandt, E., An update of the structure and 16S rRNA gene sequence-based definition of higher ranks of the class Actinobacteria, with the proposal of two new suborders and four new families and emended descriptions of the existing higher taxa, Int. J. Syst. Evol. Microbiol., 59, 589, 2009. 3. McNeil, M.M., and Brown, J.M., The medically important aerobic actinomycetes: Epidemiology and microbiology, Clin. Microbiol. Rev., 7, 357, 1994. 4. Develoux, M., Dieng, M.T., and Ndiaye, B., Mycetoma of the neck and scalp in Dakar, J. Mycol. Med., 9, 179, 1999. 5. Fahal, A.H., and Hasan, M.A., Mycetoma, Br. J. Surg., 79, 1138, 1992. 6. Trujillo, M.E., and Goodfellow, M., Numerical phonetic classification of clinically significant aerobic sporoactinomycetes and related organisms, Antonie van Leeuwenhoek, 84, 39, 2003. 7. Quintana, E.T. et al., Streptomyces sudanensis sp. nov., a new pathogen isolated from patients with actinomycetoma, Antonie van Leeuwenhoek, 93, 305, 2008. 8. Kroppenstedt, R.M., Stackebrandt, E., and Goodfellow, M., Taxonomic revision of the actinomycete genera Actinomadura and Microtetraspora, Syst. Appl. Microbiol., 13, 148, 1990. 9. Athalye, M. et al., Numerical classification of Actinomadura and Nocardiopsis, Int. J. Syst. Bacteriol., 35, 86, 1985. 10. Fischer, A., Kroppenstedt, R.M., and Stackebrandt, E., Molecular-genetic and chemotaxonomic studies on Actinomadura and Nocardiopsis, J. Gen. Microbiol., 129, 3433, 1983. 11. Poschner, J. et al., DNA–DNA reassociation and chemotaxonomic studies on Actinomadura, Microbispora, Microtetraspora, Micropolyspora and Nocardiopsis, Syst. Appl. Microbiol., 6, 264, 1985. 12. Trujillo, M.E., and Goodfellow, M., Polyphasic taxonomic study of clinically significant actinomadurae including the description of Actinomadura latina sp. nov, Zbl. Bakt., 285, 212, 1997. 13. Zhang, Z., Wang, Y., and Ruan, J., Reclassification of Thermomonospora and Microtetraspora. Int. J. Syst. Bacteriol., 48, 411, 1998. 14. Zhang, Z.S. et al., Clarification of the relationship between the members of the family Thermomonosporaceae on the basis of 16S rDNA, 16S–23S rRNA internal transcribed spacer and 23S rDNA sequences and chemotaxonomic analyses, Int. J. Syst. Evol. Microbiol., 51, 373, 2001. 15. Vincent, H., Étude sur le parasite du pied le madura, Ann. Inst. Pasteur, 8, 129, 1894. 16. Pinoy, E., Isolement et culture d’une nouvelle oospora pathogene, in Mycetome a grains rouges de la paroi thoracique, Thiroux and Pelletier (Editors), Bull. Soc. Path. Exot., 5, 585, 1912. 17. Waksman, S.A., and Henrici, A.T., Family III. Streptomy­ cetaceae Waksman and Henrici. In Breed, Murray and Hitchens (Editors), Bergey’s Manual of Determinative Bacteriology, 6th Ed., pp. 929–980, The Williams and Wilkins Co., Baltimore, 1948. 18. Becker, B., Lechevalier, M.P., and Lechevalier, H.A., Chemical composition of cell wall preparations from strains of various form genera of aerobic actinomycetes, Appl. Microbiol., 13, 236, 1965.

22 38. McIroy, J.A., Prestes, C.D., and Su, W.P., Mycetoma: Infection with tumefaction, draining sinuses, and “grains,” Cutis, 49, 107, 1992. 39. Foltz, K.D., and Fallat, L.M., Madura foot: Atypical finding and case presentation, J. Foot Ankle Surg., 43, 327, 2004. 40. Mahmoudabadi, A.Z., and Zarrin, M., Mycetomas in Iran: A review article, Mycopathologia, 165, 135, 2008. 41. Fahal, A.H., Mycetoma: A thorn in the flesh, Trans. R. Soc. Trop. Med. Hyg., 98, 3, 2004. 42. Welsh, O., Vera-Cabrera, L., and Salinas-Carmona, M.C., Mycetoma, Clin. Dermatol., 25, 195, 2007. 43. Ameen, M., and Arenas, R., Development in the management of mycetomas, Clin. Exp. Dermatol., 34, 1, 2008. 44. Gumaa, S.A., Mahgoub, E.S., and El Sid, M.A., Mycetoma of the head and neck, Am. J. Trop. Med. Hyg., 35, 596, 1986. 45. McNeil, M.M. et al., Nonmycetomic Actinomadura madurae infection in a patient with AIDS, J. Clin. Microbiol., 30, 1008, 1992. 46. Bar, W. et al., Unusual gram positive rods, causing pneumonia, Pneumologie, 57, 259, 2003. 47. Serrano, J.A. et al., The actinomycetoma in Venezuela: A  ten year study (1976–1986), Rev. Inst. Med. Trop. Sao Paulo, 30, 297, 1988. 48. Fahal, A.H., and Suliman, S.H., The clinical presentation of mycetoma, Sudan Med. J., 32, 46, 1994. 49. El Moghraby, I.M., Mycetoma in Gezira, Sudan Med. J., 9, 77, 1971. 50. Serrano, J.A. et al., Histological and ultrastructural studies on human actinomycetomas. In Szabó, Biró, and Goodfellow, M. (Editors), Biological, Biochemical and Biomedical Aspects of Actinomycetes, p. 647, Adakémiai Kiadó, Budapest, 1986. 51. Fahal, A.H., and Sabaa, A.H., Mycetoma in children in Sudan, Trans. R. Soc. Trop. Med. Hyg., 2009, Aug 27. (Epub ahead of print). 52. de Hoog, G.S. et al., Diagnostic problems with imported cases of mycetoma in The Netherlands, Mycoses, 36, 81, 1993. ‌ 53. Bapat, K.C., and Pandit, A.A., Actinomycotic mycetoma. Report of a case with diagnosis by fine needle aspiration, Acta Cytol., 35, 770, 1991.

Molecular Detection of Human Bacterial Pathogens 54. El Hag, I.A., Fahal, A.H., and Khalil, E.A.G., Fine needle aspiration of cytology of mycetoma, Acta Cytol., 40, 461, 1996. 55. Fernandes, H. et al., Cytodiagnosis of actinomycetoma, Diagn. Cytopathol., 37, 506, 2009. 56. Taha, A., A serologic survey of antibiotics to Streptomyces somaliensis and Actinomadura madurae in the Sudan using enzyme linked immunosorbent assay (ELISA), Trans. R. Soc. Trop. Med. Hyg., 77, 49, 1983. 57. Steingrube, V.A. et al., Rapid identification of clinically significant species and taxa of aerobic actinomycetes, including Actinomadura, Gordona, Nocardia, Rhodococcus, Streptomyces, and Tsukamurella isolates, by DNA amplification and restriction endonuclease analysis, J. Clin. Microbiol., 35, 817, 1997. 58. Cook, A.E., and Meyers, P.R., Rapid identification of filamentous actinomycetes to the genus level using genus specific 16S rRNA gene restriction fragment patterns, Int. J. Syst. Evol. Microbiol., 53, 1907, 2003. 59. Chun, J. et al., EzTaxon: A web-based tool for the identification of prokaryotes based on 16S ribosomal RNA gene sequences, Int. J. Syst. Evol. Microbiol., 57, 2259, 2007. 60. Welsh, O. et al., Amikacin alone and in combination with trimethoprim-sulfamethoxazole in the treatment of actinomycetomic mycetoma, J. Am. Acad. Dermatol., 17, 443, 1987. 61. Boiron, P. et al., In vitro antibiotic susceptibility testing of agents of actinomycetoma. Med. Microbiol. Lett., 1, 38, 1992. 62. Vera-Cabrera, L. et al., Therapeutic effect of a novel oxazolidinone, DA-7867, in BALB/c mice infected with Nocardia brasiliensis, PLoS Negl. Trop. Dis., 2, 1, e289, 2004. 63. ‌Steinbruge, V.A. et al., DNA amplification and restriction endonuclease análisis for differentiation of 12 Species and Taxa of Nocardia, including recognition of four new taxa within the Nocardia asteroides complex, J. Clin. Microbiol., 33, 3096, 1995. 64. Couble, A., et al., Direct detection of Nocardia spp. in clinical samples by a rapid molecular method, J. Clin. Microbiol., 43, 1921, 1998. 65. Wilson, R.W. et al., Clinical application of PCR-Restriction Enzyme Pattern Analysis for rapid identification of aerobic actinomycete isolates, J. Clin. Microbiol., 36, 148, 1998.

3 Actinomyces Debarati Paul, D. Madhusudan Reddy, Dipalok Mukherjee, Biswajit Paul, and Debosmita Paul CONTENTS 3.1 Introduction........................................................................................................................................................................ 23 3.1.1 Classification, Biology, and Epidemiology............................................................................................................. 23 3.1.2 Clinical Features and Pathogenesis........................................................................................................................ 24 3.1.3 Diagnosis................................................................................................................................................................ 25 3.1.3.1 Conventional Techniques......................................................................................................................... 25 3.1.3.2 Molecular Techniques.............................................................................................................................. 26 3.2 Methods.............................................................................................................................................................................. 26 3.2.1 Sample Preparation................................................................................................................................................. 26 3.2.2 Detection Procedures.............................................................................................................................................. 26 3.2.2.1 Single Round PCR................................................................................................................................... 26 3.2.2.2 Nested PCR.............................................................................................................................................. 27 3.2.2.3 PCR–RFLP.............................................................................................................................................. 27 3.2.2.4 Checkerboard DNA–DNA Hybridization................................................................................................ 27 3.2.2.5 Real-Time PCR for Actinomyces naeslundii and Actinomyces viscosus................................................. 28 3.3 Conclusion.......................................................................................................................................................................... 29 Acknowledgments........................................................................................................................................................................ 29 References.................................................................................................................................................................................... 29

3.1  INTRODUCTION 3.1.1  Classification, Biology, and Epidemiology The genus Actinomyces encompasses a large number of gram-positive, nonspore-forming, anaerobic, or faculta­ tively anaerobic bacterial species belonging to the family Actinomycetaceae, order Actinomycetales, phylum Actin­ obacteria, domain Bacteria. While individual Actinomyces bacteria are rods of <1 µm in diameter, Actinomyces colonies on nutrient medium are irregular in form with radiant edges,  consisting of fungus-like branched networks of hyphae. Classification and speciation of Actinomyces are based on analysis of DNA and rRNA (e.g., DNA fingerprinting and ribotyping), cell wall structure, metabolic end products, biochemical reactions, and serology. Most species represent a distinct serologic group, often with subtypes. Slack et al. were the first to apply fluorescent antibody techniques to establish and revise four serological groups within Actinomyces.1 By comparing long 16S rRNA sequences generated by reverse transcriptase, it was shown that all species formed a phylogenetically coherent cluster in which Actinomyces bovis, Actinomyces viscosus, Actinomyces naeslundii, Actinomyces odontolyticus, and Actinomyces

israelii constituted genetically well-defined species.2 To date, 36 or so Actinomyces species have been identified, of which 19 species are associated with humans and animals, with A. israelii being the most common cause of actinomycosis—a chronic suppurative and granulomatous disease of the cervico-facial, thoracic, or abdominal areas. Actinomyces are often found in healthy humans in the oral cavity, lacunas of the tonsils, or on mucosa of the gastrointestinal tract. Actinomyces, in particular A. israelii, may cause infection when they are introduced into the soft tissues by trauma, surgery, or another infection. Once in the tissues, it may form an abscess that develops into a hard red to reddish purple lump. When the abscess breaks through the skin, it forms pus-discharging lesions. Some Actinomyces spp. are opportunist pathogens associated with other mixed anaerobic infections and with infections in severely compromised patients.3 The first published clinical description of the human form of the disease appeared in 1857.4 The thoracic form was described 25 years later, but it was not until 1891 that A.  israelii, the main species responsible for the human disease, was isolated.5 Actinomycosis is endemic; sporadic cases involving humans and agricultural animals occur worldwide. According to previous surveys, the incidence of actinomycosis was about 1 per 23

24

300,000 persons in the Cleveland area in the 1970s, and about 1 per 100,000 persons in Germany and the Netherlands. A male-to-female infection ratio of 3:1 is encountered in most series. Unconfirmed explanations for this include poorer oral hygiene and augmented oral trauma in males. Cases of Actinomyces infestation in humans from sick humans or animals have not been described.

3.1.2  Clinical Features and Pathogenesis Actinomycosis (also known as Rivalta disease, big jaw, clams, lumpy jaw, or wooden tongue) is an infection commonly of the face and neck that produces abscesses (collections of pus) and open-draining sinuses (tracts in the skin) and is chiefly caused by Actinomyces israelii. The word “actinomycosis” was derived from the Greek terms “aktino,” which refers to the radiating appearance of a sulphur granule, and “mykos,” which labels the condition a mycotic disease.5 Infection can develop in individuals of all ages; however, it is rare in the pediatric population and in patients older than 60 years. Most cases are encountered in individuals in the mid decades of life. Actinomycosis is a chronic and progressive infection that does not show spontaneous remission.6 Duration of an incubation interval is not known. It can fluctuate over a wide range and reach till several years (from time of a becoming infected development of demonstrative forms of an actinomycosis). As A. israelii is a common colonizer of the vagina, colon, and mouth, its infection in host is facilitated by a breach of the mucosal barrier during various procedures (e.g., dental), aspiration, or pathologies such as diverticulitis. On a place of introduction of Actinomyces an infectious granuloma is formed that sprouts in surrounding tissues. In granulations there are abscesses that break forming fistulas. The skin lesion has secondary character. In the formation of pyeses, staphylococcal infection contamination mainly plays the secondary role. Being a chronic infectious disease, actinomycosis provokes a chronic productive and colliquative inflammatory reaction known as actinomycotic granuloma, with macroscopically visual yellowish granules produced by bacterial proliferation. The basic clinical forms of actinomycosis include (i) actinomycosis of head, tongue, and neck; (ii) a thoracic/ pulmonary actinomycosis; (iii) abdominal actinomycosis; (iv) actinomycosis of genitourinary organs; (v) skin actinomycosis; (vi) mycetoma; (vii) actinomycosis of the central excitatory system.7–9 Classically, actinomycosis occurs in cervico-facial (55%), abdominopelvic (20%), thoracic (15%), and mixed organs (10%), including skin, brain, pericardium, and extremities.10,11 Cervico-facial actinomycosis (also known as lumpy jaw) is the most common and involves both the soft and hard tissue of the head and neck region. The various infection sites may be the scalp, the paranasal sinuses, the palate, the parotid gland, the temporal bone, the tear glands, the cheeks, the lower jaw, the tongue, the larynx, and the lower pharynx.12,13 Thoracic actinomycosis occurs in thoracic lumen, or lungs. Symptoms include chest pain, fever, and weight

Molecular Detection of Human Bacterial Pathogens

loss. Primary actinomycosis of the lung and chest wall has been less frequently reported.11,14 Actinomyces is believed to invade the thoracic organ through the oropharynx, causing bronchopulmonary infection as well as pleural and chest wall involvement. The disease is usually proceeded by various forms of pulmonary infiltration or dense hilar lymphadenopathy.4,15 Accurate diagnosis is seldom made at the time of hospital admission because the disease is rather uncommon and is often unsuspected. Pathologically, thoracic actinomycosis is a parenchymal infiltrate with involvement of the chest wall often with a rib eroded.11 Due to lack of suspicion of the disease and unfamiliarity with the clinical manifestation, culture for the organisms is frequently not done at all or is incorrectly done without consideration for the microaerophilic needs of the bacterium.11 Even among experienced physicians, sometimes despite pointers to the disease, delayed diagnosis, or misdiagnosis as tuberculosis, lung abscess, or lung cancer is common,5 as it forms a mass that extends to the chest wall. Abdominal actinomycosis leads to a number of disorders that are associated with lesions in the ileocecal region. Examples frequently encountered in developed countries include colon cancer, Crohn disease, and, less commonly, infection due to Yersinia enterocolitica and Yersinia pseudo­tuberculosis. Many other infectious, neoplastic, and drug-related causes of ileocecal lesions have been described. This condition is mostly preceded by surgery such as laparotomy for acute appendicitis, perforated ulcer, or gallbladder inflammation. Infection usually begins in the gastrointestinal tract and spreads to the abdominal wall. Spiking fever and chills, intestinal colic, vomiting, and weight loss, a palpable (can be felt) mass, and an external sinus are evident in this type of actinomycosis. This type of actinomycosis may be mistaken for Crohn disease, malignancy, tuberculosis, Amebiasis (an infection of the intestine or liver), or chronic appendicitis. Genital actinomycosis and pelvic actinomycosis are rare. Actinomyces species exist in the normal flora of the gastrointestinal system and female genitourinary tracts. Abdominal actinomycosis accounts for 20% of all cases, and its diagnosis is difficult.16 Pelvic actinomycosis affects the women’s pelvic area and may cause lower abdominal pain, fever, and bleeding between menstrual periods. This form of the infection has been associated with the use of IUDs (intrauterine devices) that do not contain copper. Actinomycosis of bones and joints seems rare. The clinical presentation of spinal actinomycosis may be nonspecific and back pain is the most consistent early symptom. A case of actinomycosis involving bones and joints has been recently documented.17 The patient presented fever, pain in the neck and upper back, progressive weakness and numbness in all four limbs with difficulty ambulating, constipation, and uroschesis. Correct diagnosis is difficult because the clinical and radiological findings of actinomycosis closely resemble metastatic tumors and other infectious processes. Timely surgical debridement and decompression contributed to the prompt improvement of the patient’s conditions, and

25

Actinomyces

histopathological demonstration of the inflammatory granulation tissue and gram-positive sulfur-containing filamentous bacteria led to the correct diagnosis of actinomycosis.17 Skin actinomycosis/primary cutaneous actinomycosis is very uncommon. Subcutaneous nodules usually occur on the exposed skin, extending slowly and breaking down to form sinuses. The infection elicits a granulomatous inflammatory response in the deep dermis and subcutaneous tissue as well the underlying bone, resulting in a clinical condition called mycetoma that is characterized by the formation of grains containing aggregates of the causative organisms that may be discharged onto the skin surface through multiple sinuses. Mycetoma was first described in the mid 1800s and initially named Madura foot, after the region of Madura in India where the disease was first identified. Mycetoma caused by microaerophilic actinomycetes is termed actinomycetoma, and mycetoma caused by true fungi is called eumycetoma.18–20 Actinomycosis treatment is long term, generally with up to 1 month of intravenous penicillin G followed by weeks to months of penicillin taken orally. In addition, surgical excision and drainage of abscesses may be necessary. Many patients are cured within <6 months of antibiotic therapy. If short-term antibiotic treatment is attempted, the clinical and radiological response should be closely monitored. Cervico-facial actinomycosis is especially responsive to brief courses of antibiotic treatment.21 Table 3.1 shows some antibiotics that might be helpful in actinomycosis treatment. Conventional antibiotic therapy for actinomycosis consists of high-dose penicillin for a period of 6–12 months.21,28,29 Good results with shorter courses of antibiotic treatment have been reported, but most were preceded by extensive surgery.28,30

3.1.3  Diagnosis Actinomyces is usually part of the normal flora; its presence on sputum samples, bronchial washings, or cervicovaginal secretions may not be enough to make the diagnosis. A. israelii causes actinomycosis in which chronic granulomas

become suppurative. Tissue pus contains sulfur granules, a tangled mass of branching bacteria, which provides a diagnostic feature for actinomycosis. Diagnosis of actinomycosis usually relies on the clinical picture and the presence of sulfur granules either observed macroscopically or microscopically. Since acitinomycosis presents varied manifestations that may complicate clinical diagnosis, it is helpful to apply laboratory means for accurate detection and identification of the organism. The conventional techniques for Actinomyces identification include Gram and other stains, microscopy, microbiological culture, and serology. In recent years, the application of molecular techniques has further improved the classification and identification of members of this genus. 3.1.3.1  Conventional Techniques Actinomyces are identified as gram-positive rods that are nonacid fast in diphtheroidal arrangement. Hematoxylin-eosin staining of sulfur granules reveals basophilic masses with a radiating border of eosinophilic terminal clubs. Other useful stains used for detection are Grocott–Gomori methenaminesilver nitrate stain, periodic acid-Schiff, P-aminosalicylic acid, Goodpasture stain, and Brown–Brenn stain.18–20 For culture isolation, samples (e.g., tissue, pus, or sulfur granules) are transported on anaerobic transport media and inoculated on brain and heart infusion blood agar under anaerobic condition or enriched with carbon dioxide. Culture grows in 5–7 days and may take 2–4 weeks. A. israelii is a naerobic with clinical strains varying from obligate anaerobes to microaerophilic. Identification of Actinomyces serologically was difficult before the introduction of fluorescent antibody (FA) and geldiffusion procedures. These techniques have overcome the inherent difficulties in applying serological methods to filamentous organisms. The FA technique was first applied to Actinomyces by Slack and Moore, with the eventual establishment and revision of four serological groups by Slack and Gerencser.1 A. israeli was designated as serological group D. The species specificity of the FA technique was confirmed by

TABLE 3.1 Commonly Used Antibiotics and Their Effectiveness in Treatment of Actinomycosis MIC (mg/l) Antibiotic

MIC50

MIC90

Penicillin G Tetracycline (doxycycline) Erythromycin Clindamycin Imipenem Ceftriaxone Ciprofloxacin Metronidazole Aminoglycosides (amikacin)

0.002 0.047 0.016 0.032 NR 0.008 >32 NR NR

0.008 0.25 0.016 0.064 NR 0.094 >32 NR NR

Effectiveness Efficacious

Probable efficacy Ineffective in vitro

Reference 21, 22 21, 22 21, 23 21, 23 21, 24, 25 21, 26, 27 21, 22 21, 22 21, 23

26

Lambert et al.,31 who also reported two strains of A. israeli belonging to a second serotype. The gel-diffusion technique, first used in studying Actinomyces by King and Meyer,32 has enabled identification of A. israeli31 and elucidation of its generic relationships33,34; however, the technique is ordinarily less species specific than the FA test.31 Landfried was able to place Actinomyces cultures into serological groups corresponding to those determined by FA, but observed more cross-reactions between groups with gel-diffusion than were found with the FA test. Pirtle et al.35 reported isolation of a water-soluble, heat-stable antigen from culture supernatant fluid of A. israelii, which reacted specifically with A. israeli antiserum in gel-diffusion tests. For diagnostic purpose, the intracutaneous test with actinolysathos has some value. However, it is necessary to accept only positive and sharply positive assays as weakly positive intracutaneous tests often lead to incorrect detection. Other parameters for Actinomyces identification are based on resistance or sensitivity of growth in glycerol broth to lysozyme, urease activity, decomposition of the substrates casein, tyrosine, xanthine, and hypoxanthine. A. israelii may be identified definitively by detection with gas liquid chromatography (GLC) of acetic and lactic acid as end products of carbohydrate metabolism. 3.1.3.2  Molecular Techniques Traditional ways of identification of anaerobic gram-positive nonsporulating bacilli by isolation of the organism and studying it phenotypically by elucidation of its morphologic and biochemical characteristics and metabolic end products are associated with a need for special equipment and expertise, as well as strains that are “unidentified” because of ambiguous biochemical profiles.36 Identification by genotypic methods is particularly valuable for organisms such as Actinomyces spp., as they are poorly identified using conventional biochemical or serological methods. Some of the techniques popularly used are 16S ribosomal RNA gene sequencing, PCR–RFLP (polymerase chain reaction– restriction fragment length polymorphism)/ARDRA (amplified ribosomal DNA restriction analysis), DNA probes, and DNA–DNA hybridization. Sequence information was used to develop an oligonucleotide probe for the A. israelii serotype 1 strains, which did not react with the serotype 2 strain or with rRNA from strains of eight Actinomyces species.2 Species-specific oligonucleotide probes have been developed recently, however, due to high degree of intraspecific genetic diversity occurring within Actinomyces their utility is quite limited.2,37 On the other hand PCR–RFLP/ARDRA has proved to be a highly discriminatory, practical, and cost-effective method for detection. Smaller case studies effectively used MnlI,38–40 whereas studies with more than 500 clinical isolates used separate digestions using HaeIII and HpaI.39,41 These enzymes were not discriminatory when used alone, but when combined they were effective to differentiate all Actinomyces, Arcanobacterium, and Actinobaculum spp. to subspecies levels. These results corresponded to the conventional

Molecular Detection of Human Bacterial Pathogens

techniques as well. Only basic molecular expertise and lab facilities are required for conducting these tests.

3.2  METHODS 3.2.1  Sample Preparation Frozen cultures can be thawed and suspended in distilled water to give a cell concentration of 1–3 mg of dry weight per mL. The cell suspensions should be diluted with distilled water, and 50 µL each of the serial dilutions can be subjected directly to PCR amplification. Alternatively, 40 µL of the suspensions can be mixed with 10 µL of a proteinase K solution (1 mg/mL) and 50 µL of 40 mM Tris buffer (pH  8.0) containing 1% Tween 20, 0.2% Nonidet P-40, and 0.2% mM EDTA. This buffer is a modification of PCR buffer used for sample preparation from blood cells and other fluids. The mixture should be incubated at 60°C for 20 min and then at 95°C for 10 min. Then, the protease treated sample may be centrifuged, and 5 µL of the resultant supernatant of it used for the PCR.

3.2.2  Detection Procedures 3.2.2.1  Single Round PCR Cogulu et al.42 utilized a pair of primers (Forward: 5′-AGA GTT TGA TCC TGG CTC AG-3′ and Reverse: 5′-CCA AAA CAC CAC AAA AGT GA-3′) in a single round PCR for amplification of an A. israelii-specific 230 bp product from endodontic samplings. Procedure 1. Two sequential paper points are used to soak up the fluid in the canal and then transferred to sterile 2-mL Eppendorf tubes containing VMGA III transport medium. All samples are processed within 2 h. After thoroughly shaking the endodontic sample in a mixer for 60 s (Vortex, Scientific Industries Inc., Springfield, MA), all are frozen immediately at −20°C and stored until assayed by PCR. 2. Aliquots of each sample (1.0 mL) are centrifuged at 13,000 × g for 10 min. DNA is extracted with Genomic DNA Purification Kit (Fermentas). The DNA concentrations in clinical samples are determined by spectrophotometer measurement of the absorbance at 260 nm. 3. The PCR mixture (50 μL) is composed of 1 μM of each specific primer, 5 μL of 10× PCR buffer, 2 mM of MgCl2, 1.25 U of DNA polymerase, and 0.2-mM concentrations of dNTP. 4. DNA amplification is performed in a thermal cycler GeneAmp PCR system (Applied Biosystems) with 30 cycles of 94°C for 1 min, 60°C for 1 min, and 72°C for 2.5 min. 5. PCR amplicons are analyzed by electrophoresis in a 1.5% agarose gel at 4 V/cm in Tris-borate EDTA buffer. The gels are stained for 15 min with 0.5 μg/mL

Actinomyces

ethidium bromide and visualized under ultraviolet light. The identity of the A. israelii-specific 230 bp product is inferred by visual comparison with a molecular weight ladder (100 bp DNA ladder). 3.2.2.2  Nested PCR Hansen et al.43 described a seminested PCR targeting the 16S ribosomal RNA gene for specific detection of A. israelii in paraffin-embedded tissues. The first-round PCR utilizes oligonucleotide primers ACM-F-1 (5′-AAGTCGAACGGGTCTGCCTTG-3′, nt 5–25) and ACMR-1 (5′-TCAAAGCCTTGGCAGGCCATC-3′, nt 221–241); the nested PCR uses primers ACM-F-2 (5′-TAACCTGCC CCTCACTTCTGG-3′, nt 72–92) and ACM-R-1. First- and second-round PCR amplification is performed in the GeneAmp PCR System 9700 (Applied Biosystems) with an initial denaturation at 94°C for 5 min, followed by 30 cycles of 94°C for 30 s, 52°C for 30 s, and 72°C for 60 s. Two microliters of the first-round PCR product is used as template for the second-round PCR. Because of a broad homology of bacterial ribosomal genes, the specificity of amplification products is confirmed by direct sequencing. The results suggest a nearly complete match to the 16S ribosomal RNA gene from A. israelii but no significant homology to other species. Thus, this nested PCR protocol provides a highly sensitive method for amplification of A. israelii from nondecalcified, paraffin-embedded material. It is especially helpful in clinical cases with infected osteoradionecrosis (IORN), which do not reveal Actinomyces by means of light microscopy because the tissue specimens are small. 3.2.2.3  PCR–RFLP The oligodeoxynucleotide primers are those designed to anneal to conserved regions of the eubacterial 16S rRNA44: forward, 5′-AGAGTTTGATCCTGGCTCAG-3′ (positions 8–27 of E. coli 16S rRNA); reverse, 5′-GGTTACCTTGTTACGACT-3′ (positions 1510–1492). PCR is performed in a 100 µL volume by using a PerkinElmer Cetus thermocycler (GeneAmp PCR System 9600) and the GeneAmp PCR reagent kit with Taq polymerase (Perkin-Elmer Cetus Instruments) as specified by the manufacturer. Cycling profiles consist of 30 cycles of 94°C for 1 min, 50°C for 1 min, and 72°C for 1.5 min, followed by a postrun of 72°C for 2 min. The size of the products ~1600 bp is confirmed on 1.5% agarose gel using molecular size ladders (NEB). A restriction enzyme cocktail is used to directly speciate organisms for RFLP. The restriction enzyme cocktail (5  U enzyme) is added to approximately 1 µg of DNA (10 µL PCR product) in PCR buffer that is adjusted to contain 100 mM NaCl, 1 mM dithiothreitol, and a final concentration of 7 mM MgCl2. Restriction enzyme digests are performed at 37°C for 18 h. The reaction is halted by freezing, and restriction fragments are analyzed on 10% TBE/polyacrylamide gels or

27

3.5% agarose gel in TAE buffer run at 5V/cm. A molecular weight ladder is included in each run. The gel is stained with ethidium bromide and visualized under UV illumination. Dideoxy sequencing may be employed to further verify the identity of 16S rRNA PCR products. It is performed by using ABI kit with a cycle sequencing system as specified by the manufacturer. The modified cycle conditions are 25 cycles of 95°C for 30 s, 55°C for 52 s, and 72°C for 90 s. Automated electrophoresis and analysis of DNA sequence reactions with fluorescently labeled dideoxynucleotides are performed with a ABI Prism DNA sequencer. 3.2.2.4  Checkerboard DNA–DNA Hybridization Checkerboard DNA–DNA hybridization as described by Scoransky et al.45,46 is useful for the detection of Actino­ mycetes The protocol below is a rapid DNA–DNA hybridization method based on melting profiles in microplates (Figure 3.1).47 Whole genomic DNA probes are labeled with digoxigenin coupled to dUTP, and anti-digoxigenin antibody is utilized for detection of the hybridization reaction. Procedure 1. Genomic DNA of the reference and target strains are enzymatically digested with Sau3A, ligated to the double stranded linkers P (for the target DNA) and S (for the reference DNA). These linkers provide specific sites for PCR mediated amplification with target and reference primers P1 and S1, respectively. The oligonucleotide pairs P1 and P2 as well as S1 and S2 share complementary stretches and are designed for the formation of double stranded linkers carrying 5′-Sau3A-compatible overhangs. 2. For labeling of the PCR product, digoxygenin modified dUTP (Roche) is used in a ratio 1:19 of labeled to unlabeled dTTP. 3. Reference DNA, ligated with linker S, is amplified by PCR by applying the same conditions, except that primer S is used instead of primer P and that no labeled nucleotides are incorporated during PCR. 4. Aliquots (5 μL) of PCR products are electrophoretically separated on a 1% agarose gel and visualized by SYBR® Green (Cambrex Bio Science Rockland Inc.) staining. 5. PCR products are purified by using a QIAquick PCR purification kit (QIAGEN) and are eluted from the purification column in 50 μL sterile water. 6. The DNA of the reference strains is immobilized on MaxiSorp 96-well microplates (NalgenNunc). A total amount of 1 μg DNA in 50 μL PBS/MgCl2 (8.0 mM Na2HPO4, 1.5 mM KH2PO4, 137 mM NaCl, 2.7 mM KCl pH 7.2; supplemented with MgCl2 to a final concentration of 100 mM) is immobilized per well. Before immobilization, the DNA is denatured at 99°C for 10 min in PBS/MgCl2 buffer, immediately transferred to the microplate, and incubated at 37°C for 60 min. The solution is discarded and the microplate dried for 2 h at 60°C. The coated plates

28

Molecular Detection of Human Bacterial Pathogens

DNA from target strain

DNA from reference strain

Restriction digestion with Sau3A Attach linker P

Attach linker S

(linkers are complementary to primers in next step)

PCR using dNTPs

PCR using DIG labeled dNTPs

Immobilized on microtitre plate Used as probe for hybridization Washings Washings Anti-digoxygenin-horseradish peroxidase antibodies Washings Horseradish peroxidase (coloring agent) DNA target strain Linker S Digoxygenin label DNA reference strain Linker P Anti-digoxygenin Fab fragments

Photometric detection at 450 nm

FIGURE 3.1  Schematic representation of the DNA–DNA hybridization procedure in microplates. DNA of interest is restricted with Sau3A, and ligated with the appropriated linker (P or S). During amplification of the restricted and ligated DNAs, the DNA of the target strain is labeled with Dig dUTP. Anti-digoxygenin-HRPO antibodies are used for detection of hybridized fragments. Photometric detection of color in the wells indicates the degree of hybridization between the target and reference strain.







can be stored at 4°C for 2–3 weeks sealed with adhesive film (NalgenNunc). 7. Prior to use, the plates are washed twice with 100 μL PBS to remove excess, not immobilized DNA. The appropriate hybridization temperature (THyb) is determined applying the following formula10: THyb  = 81.5 + 16.6 × log M + 0.41 × (%G+C) – 0.7  × % formamide THyb = melting temperature [°C], M = concentration of monovalent cations). 8. Following prehybridization for 30 min at THyb with 50 μL hybridization solution [5× SSC, 1% blocking reagent (Roche), 0.1% n-laurylsarkosine, 0.02% SDS, 32% formamide], 20 ng denatured target DNA is diluted in 50 μL hybridization solution, and transferred to the coated microplates. The hybridization is performed for 2 h at THyb. The microplate cavities are washed twice for 6 min at THyb with 100 μL of the washing solutions. 9. Thermal denaturation curves are generated by applying washing solutions of increasing stringency. For the detection of DNA–DNA hybrids Commercial antidigoxygenin-horseradish peroxidase Fab fragments (anti-DIG-POD) (Roche) stock solution is used after dilution 1:1000 by adding dilution buffer [1% blocking

reagent (Roche) in PBS]. 100 μL of the antibody solution are added to each well. After incubating for 1 h at room temperature, the plates are washed three times with PBS (8.0 mM Na2HPO4, 1.5  mM KH2PO4, 137 mM NaCl, 2.7 mM KCl pH 7.2). 10. For colorimetric detection of the hybrids 100 μL of substrate solution (BM-Blue, Roche) are added to each well at room temperature. Color development is stopped after 20 min with 100 μL 1M sulfuric acid. The absorption is measured at 450 nm. 3.2.2.5  R  eal-Time PCR for Actinomyces naeslundii and Actinomyces viscosus Suzuki et al.48 described the use of TaqMan real-time PCR for detection of A. naeslundii and A. viscosus in dental plaque samples with primers derived from the 16S rRNA gene (Table 3.2). Procedure 1. Real-time PCR mixture (20-μL) is composed of 1 μL of lysed cells, 1× qPCR Master Mix (Eurogentec), forward and reverse primers at 200 nM each, and the TaqMan probe at 250 nM. Separate PCR mixtures are prepared for A. naeslundii and A. viscosus.

29

Actinomyces

TABLE 3.2 Primers and Probe for Real-Time PCR Detection of A. naeslundii and A. viscosus Primer or Probe F1 R1 Probe 1 F2 R2 Probe 2



Sequence (5′–3′) TCG AAA CTC AGC AAG TAG CCG AGA GGA GGG CCA CAA AAG AAA FAM- GGG TAC TCT AGT CCA AAC TGG CGG ATA GCG -TAMRA ATG TGG GTC TGA CCT GCT GC CAA AGT CGA TCA CGC TCC G FAM- ACG GAG GTC GGG AAC GGT GGA AG -TAMRA

2. Amplification is conducted at 50°C for 2 min and 95°C for 10 min, followed by 60 cycles of 95°C for 15 s, and 58°C for 1 min with the ABI PRISM 7700 Sequence Detection System (Applied Biosystems).



3.3  CONCLUSION



Actinomycosis is a rare disease but a serious one in case of late detection. It affects various body parts including neck/ face, lungs, alimentary canal, or the pelvic/gonadial organs. Diagnosis is based on identification of characteristic sulfur granules in whole pus, histological staining, and preferably also by a positive culture. However, molecular methods of detection are more reliable and rapid. Treatment is usually with different antibiotics and is commonly associated with surgeries depending on the extent of spread of the disease. Beginning treatment promptly helps quicken the recovery and complete recovery is possible. Development of direct detection methods may aid patient management and further elucidate clinical associations.



ACKNOWLEDGMENTS We thank Dr. Mark Lawrence and Dr. Shane Burgess for their support in writing the manuscript. We are also grateful to Dr. Rakesh K. Jain and Dr. Walter J. Diehl for their encouragement.

REFERENCES



1. Gerencser, M.A., and Slack, J.M., Serological identification of Actinomyces using fluorescent antibody techniques, J. Dent. Res., 55, A184, 1976. 2. Stackebrandt, E., and Charfreitag, O., Partial 16S rRNA primary structure of five Actinomyces species: Phylogenetic implications and development of an Actinomyces israeliispecific oligonucleotide probe, J. Gen. Microbiol., 136, 37, 1990. 3. Leek, J.H., Actinomycosis of the tympanomastoid, Laryngoscope, 84, 290–301, 1974. 4. Bennhoff, D.F., Actinomycosis: Diagnostic and therapeutic considerations and a review of 32 cases, Laryngoscope, 94, 1198, 1984. 5. Mabeza, G.F., and Macfarlane, J., Pulmonary actinomycosis, Eur. Respir. J., 21, 545, 2003. 6. Sharkawy, A.A., Cervicofacial actinomycosis and mandibular osteomyelitis, Infect. Dis. Clin. North Am., 21, 543, 2007.





Specificity A. naeslundii

A. viscosus

7. Dhaliwal, U. et al., Clinical and cytopathologic correlation in chronic inflammations of the orbit and ocular adnexa: A review of 55 cases, Orbit, 23, 219, 2004. 8. Pittman, F.E. et al., Clinical, histochemical, and electron microscopic study of colonic histiocytosis, Gut, 7, 458, 1966. 9. Richtsmeier, W.J., and Johns, M.E., Actinomycosis of the head and neck, CRC Crit. Rev. Clin. Lab. Sci., 11, 175, 1979. 10. Brown, J.R., Human actinomycosis. A study of 181 subjects, Hum. Pathol., 4, 319, 1973. 11. Hsieh, M.J. et al., Thoracic actinomycosis, Chest, 104, 366, 1993. 12. Miller, M., and Haddad, A.J. Cervicofacial actinomycosis. Oral Surg. Oral Med. Oral Pathol. Oral Radiol. Endod., 85, 496, 1998. 13. Belmont, M.J., Behar, P.M., and Wax, M.K. Atypical presentations of actinomycosis. Head Neck, 21, 264, 1999. 14. James, R., Heneghan, M.A., and Lipkansky, V., Thoracic actinomycosis, Chest, 87, 536, 1985. 15. Frank, P., and Strickland, B., Pulmonary actinomycosis, Br. J. Radiol., 47, 373, 1974. 16. Karagulle, E. et al., Abdominal actinomycosis mimicking acute appendicitis, Can. J. Surg., 51, E109, 2008. 17. Xu, G.P. et al., Cervical actinomycosis with spinal cord compression, case report and literature review, Chemotherapy, 54, 63, 2008. 18. Ramam, M. et al., A two-step schedule for the treatment of actinomycotic mycetomas. Acta Derm. Venereol., 80, 378, 2000. 19. Tendolkar, U.M. et al., Microbiological study of mycetoma patients from Bombay with special reference to actinomyces immunofluorescence, Indian J. Pathol. Microbiol., 36, 245, 1993. 20. Warren, N.G., Actinomycosis, nocardiosis, and actinomycetoma, Dermatol. Clin., 14, 85, 1996. 21. Sudhakar, S.S., and Ross, J.J., Short-term treatment of actinomycosis: Two cases and a review, Clin. Infect. Dis., 38, 444, 2004. 22. Wade, W.G., In-vitro activity of ciprofloxacin and other agents against oral bacteria, J. Antimicrob. Chemother., 24, 683, 1989. 23. Baik, J.J. et al., Pulmonary actinomycosis in Korea, Respirology, 4, 31, 1999. 24. Edelmann, M. et al., Treatment of abdominothoracic actino­ mycosis with imipenem, Eur J. Clin. Microbiol., 6, 194, 1987. 25. Yew, W.W. et al., Use of imipenem in the treatment of thoracic actinomycosis, Clin. Infect. Dis., 19, 983, 1994. 26. Skoutelis, A., Petrochilos, J., and Bassaris, H., Successful treatment of thoracic actinomycosis with ceftriaxone, Clin. Infect. Dis., 19, 161, 1994.

30 27. Rolfe, R.D., and Finegold, S.M., Comparative in vitro activity of ceftriaxone against anaerobic bacteria, Antimicrob. Agents Chemother., 22, 338, 1982. 28. Trutnovsky, G., Tamussino, K., and Reich, O., Short-term antibiotic treatment of pelvic actinomycosis, Int. J. Gynaecol. Obstet., 101, 203, 2008. 29. Fiorino, A.S., Intrauterine contraceptive device-associated actinomycotic abscess and Actinomyces detection on cervical smear, Obstet. Gynecol., 87, 142, 1996. 30. Atad, J. et al., Pelvic actinomycosis. Is long-term antibiotic therapy necessary? J. Reprod. Med., 44, 939, 1999. 31. Lambert, F.W., Jr., Brown, J.M., and Georg, L.K. Identification of Actinomyces israelii and Actinomyces naeslundii by fluorescent-antibody and agar-gel diffusion techniques. J. Bacteriol., 94, 1287, 1967. 32. King, S., and Meyer, E., Gel diffusion technique in antigenantibody reactions of Actinomyces species and “anaerobic diphtheroids,” J. Bacteriol., 85, 186,1963. 33. Slack, J.M., Landfried, S., and Gerencser, M.A., Morphological, biochemical, and serological studies on 64 strains of Actinomyces israelii, J. Bacteriol., 97, 873, 1969. 34. Snyder, M.L. et al., Studies on oral filamentous bacteria. II. Serological relationships within the genera Actinomyces, Nocardia, Bacterionema and Leptotrichia, J. Infect. Dis., 117, 341, 1967. 35. Pirtle, E.C., Rebers, P.A., and Weigel, W.W., Nitrogencontaining and carbohydrate-containing antigen from Actinomyces bovis, J. Bacteriol., 89, 880, 1965. 36. Woo, P.C. et al., Diagnosis of pelvic actinomycosis by 16S ribosomal RNA gene sequencing and its clinical significance, Diagn. Microbiol. Infect. Dis., 43, 113, 2002. 37. Jauh-Hsun, C., Vinh, T., Davies, J.K., and Figdor, D., Molecular approaches to the differentiation of Actinomyces species, Oral Microbiol. Immunol., 14, 250, 1999.

Molecular Detection of Human Bacterial Pathogens 38. Ruby, J.D. et al., Genetic characterization of the oral Actinomyces, Arch. Oral Biol., 47, 457, 2002. 39. Sato, T. et al., Differentiation of oral Actinomyces species by 16S ribosomal DNA polymerase chain reaction-restriction fragment length polymorphism, Arch. Oral Biol., 43, 247, 1998. 40. Tang, G., Samaranayake, L.P., and Yip, H.K. Genotypic diversity of oral Actinomyces naeslundii genospecies 1 and 2 in caries-active preschool children. Oral Microbiol. Immunol., 19, 371, 2004. 41. Narayanan, S. et al., Biochemical and biological characterizations and ribotyping of Actinomyces pyogenes and Actinomyces pyogenes-like organisms from liver abscesses in cattle, Vet. Microbiol., 61, 289, 1998. 42. Cogulu, D. et al., PCR-based identification of selected pathogens associated with endodontic infections in deciduous and permanent teeth, Oral Surg. Oral Med. Oral Pathol. Oral Radiol. Endodontol., 106, 443, 2008. 43. Hansen, T. et al., Actinomyces in infected osteoradionecrosis—underestimated? Hum. Pathol., 37, 61, 2006. 44. Weisburg, W.G. et al., 16S ribosomal DNA amplification for phylogenetic study, J. Bacteriol., 173, 697, 1991. 45. Colombo, A.P. et al., Clinical and microbiological features of refractory periodontitis subjects, J. Clin. Periodontol., 25, 169, 1998. 46. Socransky, S.S., Smith, C., and Haffajee, A.D., Subgingival microbial profiles in refractory periodontal disease, J. Clin. Periodontol., 29, 260, 2002. 47. Mehlen, A. et al., Development of a fast DNA–DNA hybridization method based on melting profiles in microplates, Syst. Appl. Microbiol., 27, 689, 2004. 48. Suzuki, N., Yoshida, A., and Nakano, Y. Quantitative analysis of multi-species oral biofilms by TaqMan real-time PCR, Clin. Med. Res., 3, 176, 2005.

4 Atopobium Piet Cools, Hans Verstraelen, Mario Vaneechoutte, and Rita Verhelst CONTENTS 4.1 Introduction........................................................................................................................................................................ 31 4.1.1 Characteristics of Atopobium spp........................................................................................................................... 31 4.1.2 A. vaginae and Bacterial Vaginosis........................................................................................................................ 32 4.1.3 A. vaginae in Pelvic Inflammatory Disease........................................................................................................... 33 4.1.4 Cases of Sepsis by Atopobium Species................................................................................................................... 33 4.1.5 Atopobium Species in Oral Health and Disease..................................................................................................... 33 4.2 Methods.............................................................................................................................................................................. 34 4.2.1 Phenotypic Identification........................................................................................................................................ 34 4.2.2 Genotypic Identification......................................................................................................................................... 34 4.2.2.1 Sample Preparation.................................................................................................................................. 34 4.2.2.2 Detection Procedures............................................................................................................................... 35 4.3 Future Perspectives............................................................................................................................................................. 39 References.................................................................................................................................................................................... 39

4.1  INTRODUCTION 4.1.1  Characteristics of Atopobium spp. Classification. The genus Atopobium was introduced by Collins and Wallbanks1 to reclassify species formerly designated Lactobacillus minutus (Hauduroy 1937), Lactobacillus rimae, and Streptococcus parvulus (Weinberg 1937). In 1999, Eubacterium fossor 2 and a newly described species, Atopobium vaginae,3 were placed in the genus.4 Although the genus Atopobium, belonging to the family Coriobacteraceae,5 has a low G + C content, it represents a deep branch within the high G + C gram-positive bacteria, the Actinobacteria.6 Coriobacterium glomerans is the closest phylogenetic relative.3 Today, the genus houses five species: A. vaginae (type strain CCUG 38953T), A. rimae (ATCC 49626T), A. parvulum (ATCC 33793T), A. fossor (NCTC 11919T), and A. minutum (ATCC 33267T), the latter being the type species.1 The first complete genome sequence of the genus Atopobium was published by Copeland in 2009. The A. parvulum type strain (IPP 1246T) has a 1,543,805 bp long single replicon genome with 1369 protein-coding and 49 RNA genes. Morphology. Cells consist of short elliptical rods (A.  rimae, A. minutum, A. fossor),7 with central swellings (A.  rimae)7 or small cocci (A. parvulum and A. vaginae)3,7 that may appear to be elliptical. Cells occur singly, in pairs, in clumps (A. parvulum),7 and in short chains, and are grampositive, nonmotile, and nonsporeforming.1 Colonies on

brain heart blood agar are pinpoint to 2 mm in diameter after 5 days of anaerobic incubation at 37°C. They appear translucent to transparent and are white, light yellow, or grey.1 Biology and Physiology. Atopobium species are obligately anaerobic.1 Although originally described as facultative anaerobic,3 GeissdÖrfer et al. and Ferris et al. found their A. vaginae isolates—including the type strain—to be strictly anaerobic.8,9 Atopobium species are catalase negative and produce major amounts of lactic acid from glucose, together with acetic acid and formic acids. For A. minutum, A. parvulum, and A. rimae, trace amounts of succinic acid may be detected.1,10 G + C content ranges from 39% (A. parvulum), over 44% (A. vaginae, A. fossor, and A. minutum), to 45%. Growth Requirements. Atopobium spp. are fastidious and all need to be grown under anaerobic conditions (N2 80%, H2 10%, CO2 10%) at 37°C. A. parvulum strain IPP 1246T, DSM 20469, can be grown in DSMZ medium 104 (modified PYG [http://www.dsmz. de/microorganisms/media_list.php]) or Fastidious Anaerobe Agar (FAA, LabM) supplemented with 5% horse blood. Its growth is substantially stimulated by 0.02% (vol/vol) Tween 80 and by 10% (vol/vol) rabbit serum added to culture media.7 A. vaginae can be cultured on Columbia agar with 5% horse blood3 or 5% sheep blood; tryptone soy agar supplemented with 5% sheep blood11; Brucella broth supplemented with vitamin K (1 µg/mL), hemin (5 µg/mL), and 5% laked horse blood9; Brucella agar with 5% sheep blood9 or 31

32

Chocolate agar (Nikolaitchouk, personal communication, 2010) for 2–5 days. A. fossor can be grown on DSMZ medium 110 (chopped meat medium with carbohydrates) or ATCC medium 1490 (modified chopped meat medium). A. minutum can be grown on ATCC medium 593 (chopped meat medium). A. rimae can be cultured on brain heart blood agar,7 on fastidious anaerobe agar (FAA, LabM) supplemented with 5% horse blood,12 in DSMZ medium 104, or on tryptic soy agar supplemented with 5% sheep blood.13 Growth is stimulated by 0.02% Tween 80 and by 10% rabbit serum added to culture media.7

4.1.2  A. vaginae and Bacterial Vaginosis Only recently, A. vaginae has been implicated as an important marker of bacterial vaginosis.9,13–15 Bacterial vaginosis is the single most common cause of vaginitis among women of childbearing age.16 Besides a nuisance problem, bacterial vaginosis is associated with adverse obstetric and gynecologic outcomes including premature labor, preterm birth, preterm prelabor rupture of membranes, miscarriage, intrauterine infection, postcesarean endometritis, chorioamnionitis, upper genital tract infections, and pelvic inflammatory disease.17–24 Bacterial vaginosis also renders women more vulnerable to the acquisition of sexually transmitted diseases like gonorrhea, chlamydiosis,25 genital herpes,26 and HIV-1,27 and it propagates viral replication and shedding of HIV-128 and HSV-2.29 Clinical Features. About half of all women with a diagnosis of bacterial vaginosis experience no symptoms at all. When present, characteristic symptoms are a profuse, homogeneous, off-white, and often malodorous vaginal discharge, which can cause considerable distress and discomfort. As bacterial vaginosis is rarely associated with inflammation, no inflammatory signs will be found and the discharge represents the only symptom, often accompanied with odor. Association of A. vaginae with Bacterial Vaginosis. Although controversial,30 bacterial vaginosis is typically regarded as a polymicrobial disease31 with an unknown etiology32 whereby the normal Lactobacillus dominated microflora is replaced by an overgrowth of mainly anaerobic bacteria.33,34 The bacteria most frequently isolated from women with bacterial vaginosis include Gardnerella vaginalis, A. vaginae, Mobiluncus spp., Bacteroides spp., Prevotella spp., and mycoplasmas,35,36 most of which can also be detected in women with normal microflora and by consequence are not specific for this syndrome.35 G. vaginalis is classically regarded as most associated with bacterial vaginosis, but is detected in some studies in 50%–60% of women without bacterial vaginosis.37 Recent independent studies reveal a stronger association of A. vaginae with bacterial vaginosis than G. vaginalis.9,13,14,32,38,39 This stronger association might be explained by the fact that G. vaginalis is more frequently detected in women with a normal vaginal microflora than A. vaginae, which is rarely

Molecular Detection of Human Bacterial Pathogens

(if ever) a component of a normal vaginal microflora.9,13,32,39 A.  vaginae not only appears to be more than twofold as specific for bacterial vaginosis, but its copresence with G. vaginalis was associated with significantly higher rates of recurrent bacterial vaginosis than those seen in women with only G. vaginalis.32 A. vaginae rarely occurs in the absence of G. vaginalis,32 suggesting a possible synergism between the two organisms. This suggestion was strengthened by observations of high rates of co-occurrence of A. vaginae and G. vaginalis13,39,40 and by the fact that A. vaginae was found in 50% of G. vaginalis biofilms in women with bacterial vaginosis.15 In women with preterm labor, high vaginal concentrations of A. vaginae and G. vaginalis are significantly associated with the risk of preterm delivery.24 Since many bacterial vaginosis-associated bacteria have been found within the amnion and fetal membranes of women delivering prematurely,41,42 suggesting that microbial invasion of the amniotic cavity results primarily from the ascent of bacteria from a disturbed vaginal microflora, it is worth noting that since A. vaginae has also been detected in upper reproductive tract disease,3,8,43 it may play a role in adverse pregnancy outcomes. In postmenopausal women with bacterial vaginosis, A. vaginae was found to be the predominant species.14 A. vaginae and Bacterial Vaginosis Treatment. Several studies pointed at A. vaginae as a metronidazole-resistant bacterium.8,9,32 Though we found that metronidazole susceptibility varied to a significant extent across various A. vaginae strains,11 the finding is intriguing, considering that metronidazole actually represents the standard therapy in case of bacterial vaginosis. Bradshaw et al. subsequently showed in a large prospective cohort study that rates of recurrence of bacterial vaginosis were strikingly higher among women harboring A. vaginae along with G. vaginalis, as compared to those carrying only G. vaginalis. Ferris et al. added another intriguing finding obtained from a treatment study with topical metronidazole gel. This study indicated that pretreatment A. vaginae concentrations were highest in patients who had completely or partially failed treatment, while concentrations in pretreatment A. vaginae were lowest in patients who were cured.44 In the pretreatment group, A. vaginae concentrations were highest in patients who completely or partially failed treatment, and lowest in patients who were cured. Pathogenesis. Bacterial biofilms have been associated with several recalcitrant infections like cystic fibrosis45 and chronic rhinosinusitis46 and are estimated to account for over 80% of microbial infections in the body.47 In addition, in bacterial vaginosis, the existence of a dense, adherent bacterial biofilm on the vaginal mucosa has been recently shown.48 Moreover, this biofilm, composed for more than 90% of G. vaginalis and A. vaginae, is an obligate finding in bacterial vaginosis and is absent in women with normal vaginal microflora. As a virulence mechanism, a biofilm prevents antimicrobial agents to reach bacteria—often present as latent cells within the biofilm—and enhances bacterial attachment to

Atopobium

epithelial cells, allowing higher bacterial concentrations than in luminal fluid.49 The biofilm formed in bacterial vaginosis and the synergism between G. vaginalis and A. vaginae may play a significant role in pathogenesis50 and high recurrence rates of bacterial vaginosis.15 In response to A. vaginae, vaginal epithelial cells secreted IL-6 and IL-8 in an in vitro model. Furthermore, transcription of an antimicrobial β-defensin peptide occurred after triggering the Toll-like receptor 2.51 This required live, adhering bacteria capable of protein synthesis. These data contribute indirect evidence for a putative role of A. vaginae in the pathogenesis of bacterial vaginosis.

4.1.3  A. vaginae in Pelvic Inflammatory Disease Pelvic inflammatory disease (PID) is a condition that lacks a precise definition and may include infection of any or all of the following anatomic locations: the endometrium (endometritis), the Fallopian tubes (salpingitis), the ovaries (oophoritis), the uterine wall (myometritis), the uterine serosa and broad ligaments (parametritis), and the pelvic peritoneum.52 In addition, a tuboovarian abscess may form. Some authors prefer the term salpingitis because, at a minimum, the oviduct (fallopian tube) is involved, and most of the long-term sequelae of PID result from destruction of the tubal architecture. Vaginal and endocervical infections can ascend and cause PID. Besides an indirect role for A. vaginae in the pathogenesis of PID by its putative role in pathogenesis of bacterial vaginosis, of which PID is one of the sequelae, A. vaginae has been recovered from patients with salpingitis,43 as well as from a patient with a tuboovarian abscess.8 Although the etiology of salpingitis is often attributed to sexually transmitted organisms such as Chlamydia trachomatis53 and Neisseria gonorrhoeae,54 which colonize the cervix and then ascend to the upper genital tract and cause tissue damage, the majority of cases have no known etiology.55 Recently, A. vaginae was detected in 2 of 11 Fallopian tube samples of women with salpingitis with bacterial etiology.43 A. vaginae may partly be responsible for the association between bacterial vaginosis and salpingitis.55 In 2000, a 39-year-old woman had a transvaginal oocyte recovery as part of an in vitro fertilization program. In 2003, a woman with a 3-day history of severe pain in the lower abdomen was admitted to a clinic.8 Laparotomy revealed a tuboovarian abscess. The infection spread from the abscess, causing inflammation of the peritoneum and appendix. A.  vaginae was recovered from the abscess as the sole organism, defining its clinical significance. Other isolates of A. vaginae, listed in the culture collection at the University of Göteberg in Sweden, originated from blood and amniotic membrane infections.14 A case of Atopobium vaginae causing uterine endometritis has been recently.55a

4.1.4  Cases of Sepsis by Atopobium Species In 2007, a case of A. rimae bacteremia was reported in a 47-year-old man from Seoul, Korea with liver cirrhosis.56

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The patient was admitted to the hospital due to fever and chill. Abdominal and pelvic computed tomography showed a small abscess near the left adrenal gland. In 2007, a 77-year-old woman with a history of right thoracotomy for pneumothorax was hospitalized for inhalation pneumonia caused by paralysis of the right vocal cord.57 Septic shock and a fever of 38°C developed, which was complicated by acute respiratory failure and stroke. A. rimae was identified as causative agent of the sepsis. After transferring to the intensive care unit and 7 days treatment, the patient’s condition improved and remained apyretic. In 2008, a 38-year-old male was admitted to the medical intensive care unit with sepsis.58 In addition to fever, elevated heart rate, and hypotension, physical examination revealed an extensive, necrotic decubitus ulcer of the left hip and poor oral hygiene. The Atopobium species associated with the sepsis likely originated from the ulcer. The Atopobium species reportedly showed maximum identity (99%) to Atopobium sp. oral clone C019, which has 98% sequence similarity to the type strain of A. rimae59 and was provisionally named Atopobium detroiti by the authors.

4.1.5  Atopobium Species in Oral Health and Disease Halitosis. Halitosis, or oral malodor, is a very common condition that may affect up to 30% of the population.60,61 In most cases, halitosis arises from malodorous substances—mainly volatile sulfur compounds—produced in the oral cavity as a consequence of the metabolism of bacteria colonizing the oral surfaces, periodontal pockets, and especially the dorsal tongue surface. Species typically associated with halitosis are Treponema denticola, Porphyromonas gingivalis, Porphyromonas endodontalis, and Eubacterium species,62 isolated from periodontal pockets. Whilst malodor is often associated with the presence of periodontitis, in many cases there is no such link, and evidence points to the importance of bacteria in tongue coatings that results in halitosis.60 In characterizing the tongue dorsum microflora of subjects with and without halitosis, A. parvulum was one of the species most associated with halitosis in one study,61 but with health in another.63 Further research on the role of A. parvulum in halitosis and its ability to produce volatile sulfur compounds is needed to point out a potential contribution to the clinical presentation of halitosis. One study found A. vaginae in halitosis associated tongue dorsum microflora, but with low prevalence.64 Caries. Dental caries is a process that may take place on any tooth surface where dental plaque is allowed to develop over a period of time.65 Formation of dental plaque—a biofilm—is a natural physiological process. There are three microbial hypotheses for the etiology of dental caries. The specific plaque hypothesis66 states that only a few specific species, such as Streptococcus mutans, are actively involved in the disease, the nonspecific plaque hypothesis explains caries as the outcome of the overall activity of the total plaque microflora, composed of many bacterial species,67 whereas the ecological plaque hypothesis suggests that caries is a result of a shift in the balance of the resident microflora driven by

34

changes in local environmental conditions, like acid production.68 Aas et al.69 showed that 10% of subjects with dental caries did not have detectable levels of S. mutans and that other species, among which included Atopobium spp., dominated the microflora in caries progression.69 Furthermore, in caries patients with S. mutans, other species, among which included Atopobium spp., were present at significantly higher levels than S. mutans. Keeping in mind their ability to produce high amounts of acid,1 Atopobium species are likely to play important roles in caries progression and should be added to the list of putative etiological agents of caries.69 Gingivitis. The microflora within the plaque biofilm on surfaces of teeth at the gum (gingiva) margin can cause gingivitis, which can be defined as an inflammation of the gingiva without destruction of the supporting tissues. The microflora associated with dental plaque-induced gingivitis is different than that associated with health. In a model of experimental gingivitis, A. rimae was found among the species to be positively correlated with the initiation of gingivitis, whereas A. minutum was found among the species that seemed to be present as a result of gingivitis.70–72 A clone with 98% sequence similarity to the type strain of A. rimae was isolated from a patient with acute necrotizing ulcerative gingivitis, and probably represents a new species.59 Periodontitis. Periodontitis is a bacterial driven inflammatory disease73 and can be caused by an apical extension of gingivitis. On the other hand, if pulp necrosis, which can result from caries, occurs, oral bacteria can colonize the root canal system with the formation of a biofilm.73a When the colonization progresses apically, bacteria come into direct contact with the periradicular tissues, causing apical peridontitis.74 These inflammatory reactions are short-lived and self-limiting if the colonization is transient in nature. In contrast, if bacterial contamination is persistent, immune reactions can cause destruction of surrounding tissues—periodontal ligament, cementum, and alveolar bone—leading to periradicular lesions, which can lead to tooth loss. Although also found in healthy subjects, A. minutum and A.  rimae were isolated from gingival plaque samples of patients with severe or juvenile periodontitis,75–77 and A. rimae was found in periradicular lesions.78 Furthermore, A. parvulum and A. rimae were shown to be present in infected root canals.79,80 Rather than a single organism slipping past the host defense, it is the bacterial community within the biofilm that causes periodontal tissue breakdown, classifying this disease as polymicrobial. Individual contributions of bacteria in pathogenesis of periodontitis and periodontal lesions still need to be elucidated,78 taking into consideration that A. rimae and A. parvulum are candidate pathogens.77,80,81

4.2  METHODS 4.2.1  Phenotypic Identification Commercial microbial identification systems form the backbone of microbial identification in clinical microbiology

Molecular Detection of Human Bacterial Pathogens

laboratories. They exist of a series of biochemical tests that are based on detecting bacterial enzymes, products, or metabolic activity. In principle, the combination of the enzymatic or biochemical results of the different tests enables to identify the organisms to the species level. Taken into account the fastidious nature of Atopobium species, and the fact that these tests frequently fail to identify many anaerobes, especially nonspore-forming grampositive rods,82 it is not surprising that in all cases of sepsis or abscesses involving Atopobium spp. the isolates could not be identified or were misidentified with these commercial biochemical identification systems, because Atopobium spp. are not included in the databases of these identification systems. In three cases of sepsis, A. rimae or Atopobium sp. oral clone C019 could either not be identified or were misidentified using the RapID ANA II (Remel, Lexena, KS, US), API ANA strip, or Vitek ANI card (ANI code 1600201000) identification systems (bioMérieux, Marcy-l’Etoile, France).56–58 A. vaginae isolates from a tuboovarian abscess or from patients with bacterial vaginosis were misidentified using the API ID32A (bioMérieux) or a RapID ANA II identification system as Gemella morbillorum (code 2002 0335 05).8,9

4.2.2  Genotypic Identification This overview of molecular detection and identification of Atopobium spp. focuses on nucleic acid–based techniques, with emphasis on the 16S rRNA gene. The 16S rRNA gene contains regions that are highly conserved across bacterial species—reflecting absence of evolutionary divergence—and variable regions—reflecting evolutionary divergence among species. It has proven to be a very useful phylogenetic marker employed to identify bacteria.83,84 First, we discuss the techniques that are based on the hybridization of Atopobium spp. specific DNA probes—most commonly oligonucleotide probes—targeting the variable 16S rRNA alleles. Secondly, we review Atopobium spp.specific conventional and quantitative real-time PCR (qPCR) assays, targeting the 16S rRNA gene region and the cpn60 gene. Finally, we comment on culture-dependent approaches using broad-range primers. 4.2.2.1  Sample Preparation Atopobium spp. are gram-positive bacteria, for which most extraction methods have a relatively poor recovery of nucleic acids.84 In order to detect or accurately quantify Atopobium spp. in biological samples, DNA extraction requires enzymatic pretreatment to specifically lyse the gram-positive cell wall. Mutanolysin, an enzyme that degrades the thick grampositive peptidoglycan layer, has been proven to improve DNA recovery from gram positives.85 In addition, in the pretreatment step, proteinase K can be used to break down contaminating proteins and nucleases, which are liberated during lysis of several oral bacteria.

Atopobium

4.2.2.2  Detection Procedures 4.2.2.2.1  Fluorescence In Situ Hybridization Fluorescence in situ hybridization (FISH) enables direct visualization and detection of bacterial species. A typical FISH protocol includes four steps. First, prior to the hybridization, bacteria are fixed and permeabilized; this facilitates accessibility of the fluorescent probes to the target sequence and protects the RNA from degradation by endogenous RNAses.86 Samples can be settled on membrane filters and covered with the fixing agent87, or they can be mixed with the fixing agent, and stay in solution or be transferred and dried to glass slides.88 After hybridization, a washing step removes unbound probe, and detection of the labeled bacteria is carried out by microscopy or flow cytometry.89 Probes can be labeled directly by synthesizing them with one or more modified dNTPs carrying a fluorochrome chemical side group.86 Although this is the fastest and cheapest method—the fluorescent signal can be bound to the target in a single hybridization step90 —it misses the advantage of simultaneous detection of several bacterial species provided by indirect labeling of separate probes with different fluorophores. When employing two or three dyes together, a color combination can be obtained that allows the detection of multiple targets by measuring the relative contribution of each color to the total signal.91 FISH has been used to study the composition of the microflora in human feces, including the detection of bacteria of the Atopobium cluster (i.e., belonging to the genera Atopobium, Collinsella, Coriobacterium, Cryptobacterium, and Egger­ thella).92–96 FISH has also been used for detection of this Atopobium cluster in vaginal fluids33 or in vaginal biopsies.15,48 Sample Preparation and Detection. Whole stools are collected and kept at 4°C in sterile bags for no longer than 12  h.92 The stools are kneaded mechanically at 4°C to distribute the sample evenly and 0.5 g portions are suspended in 4.5 mL PBS92 or brain heart infusion broth.93 Suspension is homogenized by vortexing with glass beads92 or by using a magnetic stirrer.93 After centrifugation, the supernatant is fixed by diluting 1:3 with 4% (wt/vol) paraformaldehyde in PBS (PFA),95 and by keeping it at 4°C for 16 h. Samples can be stored in 50% (vol/vol) ethanol-PBS at –20°C92 or –70°C.93 Vaginal biopsy specimens are fixed in nonaqueous Carnoy solution for 2 h, processed and embedded into paraffin blocks by standard techniques.15,48 4-μm thick microtome biopsy sections are placed on slides.15,48 Vaginal fluid smears on glass slides are fixed in 95% ethanol.33 The probe with sequence 5′-GGTCGGTCTCTCA ACCC-3′, targeting the 16S rRNA gene of the Atopobium cluster,15,48,92,93,95,96 is labeled with carbocyanine Cy3, carbocyanine Cy5, or fluorescein isothiocyanate (MWG Biotech, Ebersberg, Germany).15,33,48 Hybridization on glass slides is carried out at 50°C without addition of formamide92 or at 45°C with 10%–20% formamide concentrations, or in suspension at 35°C.93 To enhance visualization after hybridization, bacteria can be counterstained with 4, 6-diamidino-2-phenylindole (DAPI).15,34,48

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Detection of the fluorescent signal on the glass slides is performed with a fluorescence microscope (Nikon e600, Nikon) with a 100× oil immersion objective,15,33,48 whereas a flow cytometer (Facs Calibur, Becton Dickinson) is used for the solution.93,95 4.2.2.2.2  DNA Checkerboard Array The checkerboard DNA–DNA hybridization technique allows simultaneous analysis of several microbial species against up to 28 mixed microflora samples. It involves the extraction and labeling of total genomic DNA from a culturable microorganism for use as a probe in a hybridization assay with total genomic DNA extracted from clinical samples.97 The sample DNA is fixed in thin lanes on a membrane, and after turning the membrane 90°, up to 40 whole-genome probes are loaded into the wells and allowed to hybridize with their target sequence. In two lanes, a mixture of whole genomic DNA standards, representing a known amount of cells of each species, are fixed. The checkerboard, with the columns representing samples and the rows representing species of interest, allows a comparison of spot darkness with that of the standards, making the technique semiquantitative. This technique is being widely used for the detection of species associated with oral disease or health98–104 and for detection of several species in vaginal samples.105 For the preparation of the probes, A. vaginae, A. parvulum, and A. rimae strains are grown on Trypticase soy agar supplemented with 5% defibrinated sheep blood or Columbia agar supplemented with 5% horse blood under anaerobic conditions at 35°C–37°C.105,106 Mixtures are made with a concentration of 105 and 106 bacteria/mL in TE buffer. From these mixtures and the clinical samples, DNA is isolated using the method of Smith et al.107 The whole genome probes are labeled with digoxigenin using “DIG-High Prime” (Roche, Germany).105 The DNA samples are placed into the slots of a Minislot-30 apparatus (Immunetics, Cambridge, MA), concentrated onto a nylon membrane (Boehringer Mannheim) by vacuum, and fixed by cross-linking using UV light (Stratalinker 1800, Stratagene, La Jolla, CA), followed by baking at 120°C for 20 min.105 The membrane with fixed DNA is placed in a Miniblotter-45 apparatus (Immunetics) with the lanes of DNA at 90° to the channels of the device. Probes are loaded into the wells, and hybridization takes place at 42°C overnight. After a series of wash steps, bound probes are detected by anti-digoxigenin antibody conjugated with an alkaline phosphatase and a chemi­ fluorescent substrate (Boehringer Mannheim). Signal intensities of the samples are detected with a Lumi-ImagerTM workstation (Boehringer Mannheim) or Storm FluorImager (Molecular Dynamics, Sunnyvale, CA), and semiquantitated by comparing to the standards. 4.2.2.2.3 Reverse-Capture Checkerboard Hybridization Technology In a reverse-capture checkerboard hybridization assay, in contrast to the DNA–DNA checkerboard, the probe, rather than the sample, is first fixed to the membrane, hence the name reverse-capture checkerboard.108 Probes are made

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with a polythymidine tail, which preferentially cross-links to a membrane, leaving the probe available for hybridization. Up to 30 probes that target regions of the 16S rRNA gene can be deposited on a nylon membrane in separate horizontal lanes. The method is widely used in oral microbiology. A probe for Atopobium genomospecies C1 was included in an assay studying microbial communities associated with caries,69 whereas a probe targeting A. rimae was used in studying periodontitis81,109 and abscesses.110 DNA from caries samples is isolated using a bead beater apparatus (Biospec Products) and purified using a standard Geneclean protocol (Bio 101, Inc.).69 For periodontal samples, a protocol using proteinase K is followed.109 The 16S rRNA genes are amplified by PCR using universal primers, with the forward primer digoxigenin labeled at the 5′ end.69,109 To achieve a better performance of the PCR, particularly for samples with low numbers of bacteria, a two-step hemi-nested approach can be used.81,110 Here, using universal primers, a nearly full-length 16S rRNA gene fragment is amplified, which is used as template to run two sets of partial 16S  rRNA gene amplification, with the forward primers 5′ digoxigenin labeled, resulting in two different fragments, which together encompass the nearly full-length 16S rRNA gene. The checkerboard assay is performed with the Minislot-30 and Miniblotter-45 system (Immunetics, Cambridge, MA).81,110 Probes and standards representing 105 and 106 (and 108) cells are brought into the wells and cross-linked to a nylon membrane (Amersham-Pharmacia Biotech) by UV irradiation using a Stratalinker 1800 (Stratagene).69,81,110 The probe targeting A. rimae has the following sequence: 5′-(T)20ACGGAAGACGTATTCTCC-3′.81,110 The membrane is prehybridized at 55°C for 1 h.69 Labeled PCR products in 55°C preheated hybridization solution are denatured at 95°C for 5 min and loaded onto the membrane. Hybridization is carried out for 2 h at 55°C110 or 54°C,81 and is followed by a washing and blocking step. The membrane is incubated with anti-digoxigenin antibody conjugated with alkaline phosphatase (Roche Molecular Biochemicals) and ultrasensitive chemiluminescent substrate CDP Star is subsequently added (Roche Molecular Biochemicals). Standard chemifluorescence (Storm Technology Inc., Mountain View, CA)69 or chemifluorescence (X-ray film)81,110 is used for detection. 4.2.2.2.4  DNA Microarray DNA microarrays, first described in 1995, can be regarded as a miniaturization of the reverse-capture checkerboard approach.111 They consist of a high-density matrix of DNA probes, which are printed or synthesized on a glass or silicon slide.112 Targets typically incorporate a fluorescent label, and when applied to the array, those targets that hybridize to complementary probes are detected. Broad-range PCR amplicons of clinical DNA samples can be identified after hybridization to a microarray composed of hundreds to thousands of species-specific probes.113

Molecular Detection of Human Bacterial Pathogens

Recently, a DNA microarray named the human oral microbe identification microarray (HOMIM) (http://mim. forsyth.org/) has been developed to detect approximately 300 of the most prevalent oral species, including unculturable ones.114 The HOMIM, containing probes targeting A. parvulum, A. rimae, Atopobium sp. C019, and Atopobium sp. clone C3MLM018, is used to find bacterial associations in health and disease, to monitor effects of therapy on the oral ecology, and to perform microbial perturbation studies.114–117 For smaller samples (minimal pellet), a Ready-Lyse™ Lysozyme Solution protocol (Epicentre) is recommen­ ded for the HOMIM (http://mim.forsyth.org/homim.html). Alternatively, the QIAamp DNA Mini Kit (Qiagen) is used,115,116 or samples are placed in 50 µL TE buffer, 0.5% Tween 20 and 1 µL Proteinase K, at 55°C for 2 h, followed by 5 min at 95°C.117 Purification can be done using a MasterPure DNA Purification Kit (Epicentre). For larger samples (visible pellet), an Ultra Clean Microbial DNA Isolation Kit (MO BIO Laboratories, Inc.), using a bead-beating step, is recommended. The 16S rRNA genes are amplified in two different PCR, respectively, using forward primer 5′-CCAGAGTTTGATYMTGGC-3′ and reverse primer 5′-GAAGGAGGTGWTCCARCCGCA-3′; and forward primer 5′-GACTAGAGTTTGATYMTGGC-3′ and reverse primer 5′-GYTACCTTGTTACGACTT-3′.115,116 Alternatively, a 1:1 ratio of the above-mentioned forward primers, together with either reverse primer 5′-GYTACCTTGTTACGACTT-3′ or 5′-GAAGGAGGTGW TCCADCC-3′, can be used.117 The two PCR products are combined, purified (QIAquick PCR Purification Kit, Qiagen), labeled via the incorporation of Cy3-deoxycytidine triphosphate (GE Healthcare Biosciences) during another amplification reaction, and purified again.115–117 PCR product is mixed with hybridization buffer, heated for 5 min at 100°C, and brought on the microarray. Hybridization is performed at 55°C for 16 h in a microarray incubator (Hybex Microarray Incubation system, SciGene, Sunnyvale, CA).115,117 After washing, the microarray is scant with a high-resolution laser scanner (Axon 4000B, MDS Analytical Technologies, Sunnyvale, CA).115–117 Data from individual HOMIM signals are translated to a “bar code” format, normalized, and scored 0–5. 4.2.2.2.5 Oligonucleotide-Coupled Fluorescent Microspheres A variant of the microarray technology—where hybridization takes place on a two-dimensional surface—is a technology based on optically encoded microparticles. Pioneered by Luminex, these polystyrene microbeads are imbued with two fluorescent dyes and the ratio of these creates 100 unique spectral signatures. Each bead is coated with a different oligonucleotide probe. After PCR amplification of the targets, in which amplicons are labeled, only those beads that have a bound target will be detected by a flow cytometer. A 9-plex Luminex assay was designed for characterizing the vaginal

Atopobium

microflora and for exploring molecular diagnostics of bacterial vaginosis.118 DNA is extracted using Instagene Matrix (BioRad Laboratories, Hercules, CA). The oligonucleotide capture probes are modified by addition of a 5-amino C12 linker at the 5′ end to facilitate covalent coupling to the carboxylated fluorescent microspheres (Luminex Corporation, Austin,  TX).118 The A. vaginae probe has the following sequence: 5′-CACTCTTCATTTCTTCTACCGCAGCATT AACA-3′. The vaginal swab DNA extracts are used as templates for PCR amplification of the 5′ end of the cpn60 UT using two universal primers. These primers are modified by the addition of biotin and four phosphorothioated bases at the 5′ end of the primer. The PCR product is subsequently treated with 20 U of T7 exonuclease at room temperature for 40 min, and the single-stranded PCR product is denatured at 95°C for 10 min. Hybridization of the oligonucleotidecoupled beads to the single-stranded PCR products proceeds at 60°C for 10 min, followed by the addition of streptavidinR-phycoerythrin (Invitrogen). The mixture is analyzed on a Luminex instrument (Luminex, Austin, TX) that reports the mean fluorescence intensity (MFI) of the phycoerythrin reporter dye for each of the coupled bead types. 4.2.2.2.6  PCR Using Species-Specific Primers Conventional PCR. In conventional PCR, detection and semiquantification of the amplified sequence are performed at the end of the reaction after the last PCR cycle and involves post-PCR analysis such as size fractionation of amplicons by gel electrophoresis and image analysis. The technique is widely used for detection of A. vaginae or Atopobium spp. in studies characterizing the normal and disturbed (neo)vaginal microflora,13,14,34,37,119,120 in studying the etiology, pathogenesis, and treatment of bacterial vaginosis,121–124 as well as in exploring PCR as a molecular diagnostic tool for bacterial vaginosis.37 Quantitative PCR. Quantitative PCR (qPCR), also known as real-time PCR, is in essence the detection and quantification of a fluorescent signal directly proportional to the amount of amplicons generated by PCR. Detection is achieved by coupling a thermocycler with a fluorimeter, which measures the fluorescent signal after each amplification cycle.125 This ability to monitor the amplification reaction during its exponential phase enables users to determine the initial amount of target with great precision by comparison with the signal strength obtained with a standard dilution curve, containing well-determined amounts of target DNA. The more nucleic acid templates present at the start of the reaction, the fewer amplification cycles are required to make sufficient product that can be detected. The cycle in which a significant increase in fluorescence above the threshold is measured—referred to as the “cycle of quantification” (Cq) value—can therefore be used to calculate the quantity of DNA in the sample. Fluorescent markers can be incorporated in the amplicons by means of a intercalating fluorescent dye (SYBR® Green),

37

dual-labeled hydrolysis probes (TaqMan®), hybridization probes (Light-Cycler), molecular beacons or by scorpion probes. SybrGreen I binds in the minor groove of the DNA double helix, and fluoresces at a wavelength of 530 nm, while the fluorescence of the dye in solution is negligible.126 The main drawback of these markers is their lack of specificity, because they will bind to any double-stranded DNA sequence, nonspecific amplicons and primer dimers included. The melting temperature Tm of the amplified products can be checked using DNA melting curves at the end of the PCR to guarantee the specificity of the signal. Hydrolysis probes carry a fluorophore at the 5′ end and a quencher at the 3′ end.127 The quencher can absorb the fluorescence emitted by the fluorophore and dissipate it in the form of heat. During extension, the exonuclease activity of the DNA polymerase hydrolyses the probe and releases the fluorophore at the 5′ end. The latter, now free from its quencher, will emit a characteristic fluorescence signal that can be measured at the end of each extension period. Molecular beacons are probes with hairpin structure that, like hydrolysis probes, carry a fluorophore at the 5′ end and a quencher at the 3′ end.128 In the nonhybridized state, the probe is folded in a hairpin structure owing to the partial internal complementarity of the probe.129 In this configuration, the fluorophore and quencher are close to one another and no fluorescence is detected. During the annealing stage, the parts of the probe that form the loop anneal onto the target sequence, moving the fluorophore and its quencher apart, and the fluorescence can be detected.130 Species-specific quantitative PCR (qPCR) amplifies the same diagnostic amplicons as in conventional PCR, but in contrast to conventional PCR, amplification is followed during each replication cycle. qPCR has been widely used for the detection of A. vaginae or Atopobium spp. in studies characterizing the normal and disturbed vaginal microflora,39,131,132 and in studying the etiology, pathogenesis, sequelae, treatment, and diagnosis of bacterial vaginosis.24,32,130,131,133–135 qPCR is also used for detection of the Atopobium cluster in studies characterizing the microflora in human feces.94,136 Sample Preparation and Genomic DNA Extraction. Fecal samples (20 mg) are washed three times by suspending them in 1 mL PBS and centrifuging each preparation at 14,000 × g to remove possible PCR inhibitors.94 Vaginal swabs samples are typically agitated or vortexed in 1 mL PBS14,118 or 2 mL saline37,121 to dislodge and wash the cells, followed by pelleting and resuspending the cells. Alternatively, swabs are vortexed in 400 µL lysis buffer.13,39,120,122 For DNA extractions, commercial kits or (semi) automated DNA extraction platforms are most often used, these include the DNeasy Tissue Kit (Qiagen),132 AquaPure genomic DNA kit (Bio-Rad),123 Wizard DNA purification kit (Promega),137,138 QIAamp DNA Mini Kit (Qiagen)38,135 with minor modifications involving mutanolysin pretreatment,13,39 UltraClean Soil DNA Isolation kit,37,121,133,134 QIAamp stool kit,33 Instagene matrix (Bio-Rad Laboratories, Hercules, CA),14,118 Automated

38

Molecular Detection of Human Bacterial Pathogens

MagNA Pure LC with the DNA Isolation Kit I protocol (Roche Diagnostics),32,131 MasterPure DNA purification kit (Epicentre),124 the automated X-tractor Gene system (Corbett Robotics, Brisbane, Australia),130 or the NucliSens EasyMAG platform (bioMérieux).120 A DNA extraction method for fecal samples is described by Matsuki et al.94 Primers and Controls. The primers and probes applicable for amplification and detection of Atopobium spp. are listed in Table 4.1. Composition of the PCR mixtures and cycling conditions can be found in the accompanying references. The β-globine gene,32,37,121,131,133 a human albumin gene,24,135 or the human 18S rRNA gene134 can be used as an internal control to assure that amplifiable DNA was successfully extracted and to monitor for PCR inhibitors. Alternatively, amplification of exogenous DNA from a jellyfish gene can be used to test for PCR inhibition, defined as a > two-cycle increase in the Cq for the extracted vaginal DNA sample compared to the no-template controls.133,134 Detection. After separation on 2% agarose gels, conventional PCR products are typically visualized by staining 5 µL of amplified product with ethidium bromide and placing the

gel under UV light.13,14,37,119–122,124 Specificity is confirmed by detection of the appropriate sized PCR product,124 which can be sequenced as an additional control.14,33,37 For detection in qPCR assays, the LightCycler (Roche),132 Stratagene MX 3000P,24,135 RotorGene 3000,130 and ABI platforms (Applied Biosystems) 39,94,118,133,134,136 have been used. In assays using SybrGreen I, analysis of the melting curve can be carried out after amplification to distinguish the targeted PCR product from aspecific nontargeted PCR products. For quantification, a standard curve is made. DNA, extracted from A. vaginae cultured on TSA + 5% sheep blood (Becton Dickinson) and diluted in a 1/10 series, can be used as standards.39,132 Alternatively, a serial diluted suspension of cloned 16S rRNA24,133,135 or cpn60118 plasmids can be used. 4.2.2.2.7  Sequence-Based Identification In cases were A. rimae, A. vaginae, or Atopobium sp. oral clone C019 isolates could not be identified or were misidentified using conventional techniques, the 16S rRNA gene was (partially) sequenced to reach identification.8,13,56–58

TABLE 4.1 Primers and Probes Used for Amplification and Detection of Atopobium spp. Primer and Probe Sequences (5′–3′)

Target Sequence

Target Species

Reference

GGTGAAGCAGTGGAAACACT ATTCGCTTCTGCTCGCGCA

16S rRNA gene

A. vaginae

39

CCCTATCCGCTCCTGATACC CCAAATATCTGCGCATTTCA VIC-GCAGGCTTGAGTCTGGTAGGGGA-TAMRA

16S rRNA gene

A. vaginae

24,135

GGTCGGTCTCTCAACCCGG TCATGGCCCAGAAGACCGCC FAM-CAGATTTAACTCCTGACCTAACAGACC-TAMRA

16S rRNA gene

A. vaginae

32,131

GCGAATATGGGAAAGCTCCG GAGCGGATAGGGGTTGAGC

16S rRNA gene

A. vaginae

120,122

TAGGTCAGGAGTTAAATCTG TCATGGCCCAGAAGACCGCC

16S rRNA gene

A. vaginae

123,124

Cpn60

A. vaginae

118

CAGCGTTCCTGTTACTCC GCCACTTGAAGGTCTTGC cgcgatcATTCCACGGTTGCACATAACATGCgatcgcga

16S rRNA gene

A. vaginae

130

GGTCGGTCTCTCAACCC CTCCTGACCTAACAGACC

16S rRNA gene

A. vaginae

14

GCAGGGACGAGGCCGCAA GTGTTTCCACTGCTTCACCTAA

16S rRNA gene

Atopobium spp.

37,121

TAGGCGGTYTGTTAGGTCAGGA CCTACCAGACTCAAGCCTGC FAM-CTCAACCCCTATCCGCTCCTGAT-TAMRA

16S rRNA gene

Atopobium spp.b

133,134

GGGTTGAGAGACCGACC CGGRGCTTCTTCTGCAGG

16S rRNA gene

Atopobium clusterc

94,132

CTTTGCAACCAACAATGACACC CAGCGCATACGGTAAGCGTAC

a b c

Uppercase letters indicate homology to the target sequence. Atopobium spp.: Atopobium sp. AY738657, AY738658. Atopobium cluster: Members of the genera Atopobium, Collinsella, Eggerthella, and Coriobacterium.

Atopobium

39

TABLE 4.2 Primers Used for Amplification of the 16S rRNA Gene Primer

Sequence 5′–3′

Amplicon Size

Reference

fD1 rP2

AGAGTTTGATCCTGGCTCAG ACGGCTACCTTGTTACGACTT

1454 bp

57

8FPL 806R

AGTTTGATCCTGGCTCAG GGACTACCAGGGTATCTAAT

798 bp

56

10f 534r

AGTTTGATCCTGGCTCAG ATTACCGCGGCTGCTGG

524 bp

13

515FPL 13B

TGCCAGCAGCCGCGGTAA AGGCCCGGGAACGTATTCAC

785 bp

56

Anaerobic blood cultures or swabs can be (sub)cultured on a rich blood agar (e.g., Columbia agar with 5% horse blood3) and incubated anaerobically for 2–5 days at 37°C (see growth requirements). For DNA extraction, commercial kits such as the QIAamp tissue kit (QIAGEN)57 or GenElute Bacterial Genomic DNA kit56 can be used, preferentially after pretreatment with mutanolysin. Alternatively, DNA is extracted from a colony by simple alkaline extraction.139 For sequencing purposes, 16S rRNA PCR amplification can be performed using 16S rRNA universal primers, as listed in Table 4.2. To obtain pure amplicons, excess primers and nucleotides are removed using a commercial kit (QIAquick PCR Purification Kit, Qiagen).13 Thereafter, the 16S rRNA gene amplicons are sequenced, either partially58 or fully.8,57,58 Commercial packages such as the MicroSeq® Full Gene or 500 16S rRNA Bacterial Identification Kit (PE Applied Biosystems) can be used.58 For identification, the sequence is compared to a public database (e.g., Genbank) or a curated database (e.g., SmartGene IDNS Bacteria) by using analysis software (BLAST version 2.2.957 or SmartGene [www.idnssmartgene.com]).

4.3  FUTURE PERSPECTIVES Highly sensitive high throughput analysis through deep pyrosequencing will further enlighten our knowledge of bacterial diversity and ecological dynamics associated with oral, reproductive tract, and intestinal tract infection(s). Knowledge obtained by this metagenomic approach might then be translated into rapid diagnostic assays for profiling a selected set of microbial communities in a lab-based setting. In case of bacterial vaginosis, distinct vaginal microflora patterns may be found in both intermediate and bacterial vaginosis microflora, thereby possibly differentiating intermediate microflora regressing to normal or developing to bacterial vaginosis and differentiating bacterial vaginosis microflora with different pathologic sequelae. Furthermore, molecular diagnosis of bacterial vaginosis, with a central role for A. vaginae, is now pending.37,118,133,135 Species-specific quantitative PCR techniques are being patented and will become commercially available as diagnostic assays. Also, ongoing sequence analysis of the whole genome

of A. vaginae within the A. vaginae Genome Project (http:// med.stanford.edu/sgtc/research/atopobium_vaginae.html) will probably reveal as yet undescribed virulence factors and will also improve our understanding of the pathogenic role of A. vaginae in reproductive tract infections. To explore the role of A. parvulum and A. rimae in oral health and disease, microarrays like the TNO oral microarray (O-chip, TNO, Zeist, The Netherlands) or the HOMIM (http://mim.forsyth.org/homim.html) offer the possibility to detect about 300 oral species. In cases of bacteremia, even if Atopobium spp. are included in future commercial biochemical tests, sequencing the 16S rRNA gene or the cpn60 gene will remain the golden standard to identify these species, as they are generally slow growing and biochemically nonreactive.12

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42 92. Harmsen, H.J. et al., Development of 16S rRNA-based probes for the Coriobacterium group and the Atopobium cluster and their application for enumeration of Coriobacteriaceae in human feces from volunteers of different age groups, Appl. Environ. Microbiol., 66, 4523, 2000. 93. Rigottier-Gois, L. et al., Fluorescent hybridisation combined with flow cytometry and hybridisation of total RNA to analyse the composition of microbial communities in human faeces using 16S rRNA probes, FEMS Microbiol. Ecol., 43, 237, 2003. 94. Matsuki, T. et al., Use of 16S rRNA gene-targeted groupspecific primers for real-time PCR analysis of predominant bacteria in human feces, Appl. Environ. Microbiol., 70, 7220, 2004. 95. Mah, K.W. et al., Effect of a milk formula containing probiotics on the fecal microbiota of asian infants at risk of atopic diseases, Pediatr. Res., 62, 674, 2007. 96. Kleessen, B. et al., Microbial and immunological responses relative to high-altitude exposure in mountaineers, Med. Sci. Sports Exerc., 37, 1313, 2005. 97. Socransky, S.S. et al., “Checkerboard” DNA–DNA hybridization, Biotechniques, 17, 788, 1994. 98. Nascimento, C. et al., DNA checkerboard method for bacterial pathogen identification in oral diseases, Int. J. Morphol., 24, 619, 2006. 99. Ximenez-Fyvie, L.A., Haffajee, A.D., and Socransky, S.S., Microbial composition of supra- and subgingival plaque in subjects with adult periodontitis, J. Clin. Periodontol., 27, 722, 2000. 100. Siqueira, J.F. et al., Endodontic infections: Concepts, paradigms, and perspectives, Oral Surg. Oral Med. Oral Pathol. Oral Radiol. Endod., 94, 281, 2002. 101. Socransky, S.S. et al., Microbial complexes in subgingival plaque, J. Clin. Periodontol., 25, 134, 1998. 102. Sakamoto, M., Umeda, M., and Benno, Y., Molecular analysis of human oral microbiota, J. Periodont. Res., 40, 277, 2005. 103. Colombo, A.P. et al., Clinical and microbiological features of refractory periodontitis subjects, J. Clin. Periodontol., 25, 169, 1998. 104. Haffajee, A.D. et al., Subgingival microbiota in healthy, well-maintained elder and periodontitis subjects, J. Clin. Periodontol., 25, 346, 1998. 105. Nikolaitchouk, N. et al., The lower genital tract microbiota in relation to cytokine-, SLPI- and endotoxin levels: Application of checkerboard DNA–DNA hybridization (CDH), APMIS, 116, 263, 2008. 106. Brito, L.C. et al., Use of multiple-displacement amplification and checkerboard DNA–DNA hybridization to examine the microbiota of endodontic infections, J. Clin. Microbiol., 45, 3039, 2007. 107. Smith, G.L., Socransky, S.S., and Smith, C.M., Rapid method for the purification of DNA from subgingival microorganisms, Oral Microbiol. Immunol., 4, 47, 1989. 108. Paster, B.J., Bartoszyk, I.M., and Dewhirst, F., Identification of oral streptococci using PCR-based, reverse-capture, checkerboard hybridization, Methods Cell Sci., 20, 223, 1998. 109. Paster, B.J. et al., Bacterial diversity in necrotizing ulcerative periodontitis in HIV-positive subjects, Ann. Periodontol., 7, 8, 2002. 110. Siqueira, J.F., Jr., and Rocas, I.N., The microbiota of acute apical abscesses, J. Dent. Res., 88, 61, 2009. 111. Schena, M. et al., Parallel human genome analysis: Microarraybased expression monitoring of 1000 genes, Proc. Natl. Acad. Sci. USA, 93, 10614, 1996.

Molecular Detection of Human Bacterial Pathogens 112. Mothershed, E.A., and Whitney, A.M., Nucleic acid-based methods for the detection of bacterial pathogens: Present and future considerations for the clinical laboratory, Clin. Chim. Acta 363, 206, 2006. 113. Palmer, C. et al., Rapid quantitative profiling of complex microbial populations, Nucleic Acids Res., 34, 2006. 114. Olsen, I. et al., Cultivated and not-yet-cultivated bacteria in oral biofilms, Microb. Ecol. Health Dis., 65, 2009. 115. Preza, D. et al., Microarray analysis of the microflora of root caries in elderly, Eur. J. Clin. Microbiol. Infect. Dis., 28, 509, 2009. 116. Preza, D. et al., Diversity and site-specificity of the oral microflora in the elderly, Eur. J. Clin. Microbiol. Infect. Dis., 28, 1033, 2009. 117. Colombo, A.P. et al., Comparisons of subgingival microbial profiles of refractory periodontitis, severe periodontitis, and periodontal health using the human oral microbe identification microarray, J. Periodontol., 80, 1421, 2009. 118. Dumonceaux, T.J. et al., Multiplex detection of bacteria associated with normal microbiota and with bacterial vaginosis in vaginal swabs by use of oligonucleotide-coupled fluorescent microspheres, J. Clin. Microbiol., 47, 4067, 2009. 119. Burton, J.P. et al., A preliminary survey of Atopobium vaginae in women attending the Dunedin gynaecology out-patients clinic: Is the contribution of the hard-to-culture microbiota overlooked in gynaecological disorders? Aust. N. Z. J. Obstet. Gynaecol., 45, 450, 2005. 120. Weyers, S. et al., Microflora of the penile skin-lined neovagina of transsexual women, BMC Microbiol., 9, 102, 2009. 121. Marrazzo, J.M. et al., Relationship of specific vaginal bacteria and bacterial vaginosis treatment failure in women who have sex with women, Ann. Intern. Med., 149, 20, 2008. 122. Verstraelen, H. et al., Gene polymorphisms of Toll-like and related recognition receptors in relation to the vaginal carriage of Gardnerella vaginalis and Atopobium vaginae, J. Reprod. Immunol., 79, 163, 2009. 123. Ferris, M.J., Masztal, A., and Martin, D.H., Use of species-directed 16S rRNA gene PCR primers for detection of Atopobium vaginae in patients with bacterial vaginosis, J. Clin. Microbiol., 42, 5892, 2004. 124. Haggerty, C.L. et al., Clinical characteristics of bacterial vaginosis among women testing positive for fastidious bacteria, Sex. Transm. Infect., 85, 242, 2009. 125. Higuchi, R. et al., Kinetic PCR analysis: Real-time monitoring of DNA amplification reactions, Biotechnology (N Y), 11, 1026, 1993. 126. Arya, M. et al., Basic principles of real-time quantitative PCR, Expert. Rev. Mol. Diagn., 5, 209, 2005. 127. Koch, W.H. et al., Technology platforms for pharmacogenomic diagnostic assays, Nat. Rev. Drug. Discov., 3, 749, 2004. 128. Kim, Y., Sohn, D., and Tan, W., Molecular beacons in biomedical detection and clinical diagnosis, Int. J. Clin. Exp. Pathol., 1, 105, 2008. 129. Abravaya, K. et al., Molecular beacons as diagnostic tools: Technology and applications, Clin. Chem. Lab. Med., 41, 468, 2003. 130. Trama, J.P. et al., Rapid detection of Atopobium vaginae and association with organisms implicated in bacterial vaginosis, Mol. Cell. Probes, 22, 96, 2008. 131. Tabrizi, S.N. et al., Prevalence of Gardnerella vaginalis and Atopobium vaginae in virginal women, Sex. Transm. Dis., 33, 663, 2006.

Atopobium 132. Biagi, E. et al., Quantitative variations in the vaginal bacterial population associated with asymptomatic infections: A realtime polymerase chain reaction study, Eur. J. Clin. Microbiol. Infect. Dis., 28, 281, 2009. 133. Fredricks, D.N. et al., Changes in vaginal bacterial concentrations with intravaginal metronidazole therapy for bacterial vaginosis as assessed by quantitative PCR, J. Clin. Microbiol., 47, 721, 2009. 134. Mitchell, C.M et al., Comparison of oral and vaginal metronidazole for treatment of bacterial vaginosis in pregnancy: Impact on fastidious bacteria, BMC Infect. Dis., 9, 89, 2009. 135. Menard, J.P. et al., Molecular quantification of Gardnerella vaginalis and Atopobium vaginae loads to predict bacterial vaginosis, Clin. Infect. Dis., 47, 33, 2008.

43 136. Matsuda, K. et al., Establishment of an analytical system for the human fecal microbiota, based on reverse transcriptionquantitative PCR targeting of multicopy rRNA molecules, Appl. Environ. Microbiol., 75, 1961, 2009. 137. Yamamoto, T. et al., Bacterial populations in the vaginas of healthy adolescent women, J. Pediatr. Adolesc. Gynecol., 22, 11, 2009. 138. Zhou, X. et al., Characterization of vaginal microbial communities in adult healthy women using cultivation-independent methods, Microbiol. Sgm., 150, 2565, 2004. 139. Vaneechoutte, M. et al., Isolation of Moraxella canis from an ulcerated metastatic lymph node, J. Clin. Microbiol., 38, 3870, 2000.

5 Bifidobacterium Abelardo Margolles, Patricia Ruas-Madiedo, Clara G. de los Reyes-Gavilán, Borja Sánchez, and Miguel Gueimonde CONTENTS 5.1 Introduction........................................................................................................................................................................ 45 5.1.1 Classification and Taxonomy.................................................................................................................................. 45 5.1.2 Biology and Physiology.......................................................................................................................................... 46 5.1.3 Clinical Features and Pathogenesis........................................................................................................................ 49 5.1.4 Diagnosis................................................................................................................................................................ 51 5.1.4.1 Phenotypic Techniques............................................................................................................................ 51 5.1.4.2 Molecular Techniques.............................................................................................................................. 52 5.2 Methods.............................................................................................................................................................................. 53 5.2.1 Sample Preparation................................................................................................................................................. 53 5.2.2 Detection Procedures.............................................................................................................................................. 53 5.3 Conclusion and Future Perspectives................................................................................................................................... 54 References.................................................................................................................................................................................... 54

5.1  INTRODUCTION 5.1.1  Classification and Taxonomy The genus Bifidobacterium includes high G + C gram-­positive nonspore-forming, nonmotile, and nonfilamentous polymorphic rod-shaped bacteria that can display slight bends or a large variety of branchings, the slightly bifurcated club-shaped or spatulated extremities being the most commonly found. They can be organized singly or in chains, in star-like aggregates, in “V,” or in palisade arrangements when grown in vitro. They are strictly anaerobic, although some species, such as Bifidobacterium animalis and Bifidobacterium psychraerophilum, can tolerate moderately high oxygen concentrations, and they have a fermentative metabolism.1,2 Bifidobacteria were first described at the beginning of the twentieth century and were included among the family Lactobacillaceae.3 In 1924, Lactobacillus bifidum was reclassified as the new genus Bifidobacterium by Orla-Jensen.1 Currently, over 30 species are included in the genus Bifidobacterium. They form a close phylogenetic group and show high similarity (>93%) in the 16S rRNA sequences.4 The Bifidobacterium genus is included in the phylum Actino­ bacteria, class Actinobacteria, subclass Actinobacteridae, order Bifidobacteriales, and family Bifido­bacteriaceae. Phylogenetic studies, based on sequence comparison of 16S rRNA sequences and housekeeping genes, have clustered some Bifidobacterium species in six phylogenetic clusters named Bifidobacterium boum, Bifidobacterium

asteroides, Bifidobacterium adolescentis, Bifidobacterium longum, Bifidobacterium pullorum, and Bifidobacterium pseudolongum.5 Other species have displayed a large phylogenetic distance and have not been assigned to any specific cluster (Figure 5.1). The species included in the genus Bifidobacterium are B. adolescentis, B. angulatum, B. animalis, B.  asteroides, B.  bifidum, B.  bombi, B.  boum, B.  breve,  B. catenulatum, B.  choerinum, B. coryneforme, B. cuniculi, B. dentium, B. gallicum, B. gallinarum, B. indicum, B.  longum, B.  magnum, B.  merycicum, B.  minimum, B.  mongoliense, B.  pseudocatenulatum, B.  pseudolongum, B. psychraerophilum, B. pullorum, B. ruminantium, B. saeculare, B. scardovii, B. subtile, B. thermacidophilum, B. thermophilum, and B. tsurumiense.6–9 Further­more, B. animalis comprises of two subspecies (animalis and lactis), as well as B. pseudolongum (subsp. globosum and pseudolongum), and B. thermacidophilum (subsp. thermoacidophilum and porcinum), whereas the species B.  longum is subdivided in the subspecies longum, infantis, and suis (Figure 5.1). Bifidobacterium species can be isolated from a limited number of habitats—that is, human and animal gastrointestinal tract (GIT), food, insect intestine, and sewage.10,11 Among the species most commonly found in human intestine and feces are B. catenulatum, B. pseudocatenulatum, B. adolescentis, B.  longum, B. pseudolongum, B. breve, B. angulatum, B. bifidum, and B. dentium, whereas B. animalis is the species more often found in functional foods.12 In this respect, it is of key importance to remark that a clear 45

46

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D86183 B. dentium

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B.

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B.

B. adolescentis group

1 874 M5 um inim B. m 8 932 78 D8 93 are D8 cul tile sae B. sub

B.

B. pullorum group

B. adolescentis NC008618 B . ru min anti me um D B. ryc 8619 icu an 7 mD gu 861 la 92 tu m D8 61 82

Molecular Detection of Human Bacterial Pathogens

tum ula 7 ten a m D8618 c . tenulatu

BB . pseudoca

B . scard ovii AJ3 07005

24 S836 ifdum b . B

B. longum group

B. boum group

U10151 ophilum B . therm

B. breve AB006658

s . infanti m subsp 86184 B . longu D s i u s sp. 3 sub 874 gum 9 um M5 lon 873 g n . 5 lo p . s B b M su m gu n lo B.

B. thermacidophilum subsp. thermacidop hilum

AY 14 84 70

B. asteroides group

58734 iculi M B . cun

B. magnum M58740

B . gall icum D 86189

88 61 D8 m cu 30 di 587 in sM B. 49 ide ero 1275 ast i EU B. omb B. b 4110 se AB2 umien B . tsur

B . B . bo um D AB016246 the rm 8619 aci 0 do ph ilu m B. c sub oryn sp. efor po me rci M5 nu 873 m 3

5 18 86 .D bsp s su 3 ali X8951 im . lactis an s subsp B . B . animali

B . p seud B. olon pse gum ud subs B o p . l g o . lo ngu bosu ch mD m oe 8619 sub rin 4 sp u . p m seu do lon g u m D8 61 95

D8 61 86

0.005

082 432 AF

B. pseudolongum group FIGURE 5.1  Evolutionary relationships of bifidobacteria.

ecological specialization seems to exist for different bifidobacterial taxa, and species distribution depends on host age and localization in the host (i.e., intestinal mucosa versus luminal/fecal environment).13,14 Furthermore, novel metagenomic studies have analyzed the bifidobacterial microbiota in the human gut and revealed that its biodiversity was much broader than had been previously expected,14,15 showing the existence of uncharacterized phylotypes that have passed unnoticed until recently, and highlighting the importance of novel culture-independent approaches to complement the more traditional culture-dependent techniques.

5.1.2  Biology and Physiology As stated above, the natural habitat of the genus Bifidobac­ terium is the GIT of mammals, birds, and insects. The human GIT hosts a very complex microbiota that plays an important role on the health and wellbeing of the host.16 This microbial ­community reaches one of the highest cell densities recorded for any ecosystem, although its taxonomical diversity is one of the lowest. A small number of taxa dominate this ecosystem in adult humans, mainly representatives of Bacteroidetes and Firmicutes divisions.17 The phylum Actinobacteria, including the Bifidobacteriaceae family, is also present.

Bifidobacterium

The GIT colonization of newborn babies starts at birth and increases in number and diversity in few days, bifidobacteria being one of the first and most abundant colonizers. Several factors affect this process, such as the type of delivery, the initial way of feeding (breast or infant formula), or the geographical region, among others. The number of bifidobacteria declines with the age of the host, remaining more or less stable during the adult life and tending to decrease in the elderly.18 In addition, intersubject species variability in bifidobacterial populations has been noticed, but low diversity is found between different GIT regions of the same subject, denoting scarce intrasubject variability.14 At the beginning of the twentieth century, Tissier3 described for the first time bifidobacterial strains (originally named Bacillus bifidus) isolated from breastfed infant feces. During recent years, an increasing number of articles have been published reporting on the bifidobacterial diversity of the GIT ecosystem. Commonly, these studies are based on fecal sampling, leading to the isolation of new strains. However, in the fecal material, transitory bacteria derived from the diet are usually present, and therefore not representing the mucosaadhered microbiota.19 For this reason, different GIT sections, such as colonic mucosa obtained from biopsies of healthy individuals, are also used to explore the bifidobacterial diversity of the human microbiota.15 In addition, Bifidobacterium strains can also be isolated from human saliva20 and breast milk,21 as well as from sewage, in which bifidobacteria are used as indicators of fecal contamination.22 Regarding the physiology of this genus, the optimal growth temperature of most human isolated strains is about 36°C– 38°C, being higher for that of animal origin (41°C–43°C). Viability of bifidobacteria at acidic pH values is very variable but, in general, it can be considered that resistance of bifidobacteria to acidic pH is weak, with the exception of B. animalis.23 This species satisfactorily tolerates acidic pH and, in addition, the presence of oxygen, for these reasons being the most used Bifidobacterium spp. in fermented probiotic products. It has been suggested that the acid tolerance in Bifidobacterium is linked to the activity of the membrane-bound F0F1-ATPase, an enzyme responsible for the maintenance of the intracellular pH homeostasis in anaerobic bacteria through the ATPdependent extrusion of protons.24 The high F0F1-ATPase activity induction observed for this species appears to be responsible for its elevated acid resistance, this activity not being induced in nonresistant strains.25 Tolerance to bile salts is another crucial property for the persistence of bifidobacteria in the human GIT. Usually, deconjugated bile salts are more toxic than their corresponding taurine or glycine conjugates. The enzyme responsible for bile salts deconjugation is the bile salt hydrolase (BSH), but to date there is no solid evidence of a relationship between this enzyme and the levels of resistance to bile in bifidobacteria.26 Bifidobacteria have a specific pathway for the metabolism of sugars, the fructose-6-phosphate phosphoketolase (F6PPK) pathway or “bifido shunt” (Figure 5.2; Table 5.1), described for the first time in the work of Scardovi and Trovatelli.27 The key enzyme of this pathway is a xylulose-

47

5-phosphate/fructose-6-phosphate phosphoketolase (Xfp), which catalyzes the phosphorolytic cleavage of fructose-6phosphate and/or xylulose-5-phosphate. To date, two types of Xfp activities have been described in prokaryotes and eukaryotes—fructose-6-phosphate phosphoketolase (F6PPK, EC 4.1.2.22), which catalyzes the conversion of fructose-6phosphate to erythrose-4-phosphate and acetyl-phosphate; and xylulose-5-phosphate phosphoketolase (EC 4.1.2.9), which cleaves xylulose-5-phosphate, rendering acetyl-phosphate and glyceraldehyde-3-phosphate.28 In bifidobacteria, in addition to the F6PPK, a dual xylulose-5-phosphate/fructose6-phosphate phosphoketolase able to act over both fructose6-phosphate and xylulose-5-phosphate has been described.29 Bifidobacteria also present a transaldolase (Tal, EC 2.2.1.2) and a transketolase (Tkt, EC 2.2.1.1) able to convert the fructose-6-phosphate in acetyl-phosphate and glyceraldehyde-3phosphate. An acetate kinase (AckA, EC 2.7.2.1) allows the conversion of acetyl-phosphate into acetate with the concomitant production of ATP. The glyceraldehyde-3-phosphate, through the last reactions of the Embden–Meyerhof–Parnas (EMP) pathway, is metabolized to pyruvate and the activity of lactate dehydrogenase NAD-dependent (Ldh2, EC 1.1.1.27) converts this intermediate metabolite in lactate. In addition, pyruvate in combination with Co-A is converted into formate and acetyl-CoA by the activity of the formate C-acetyltransferase (Pfl, EC 2.3.1.54). Finally, acetyl-CoA is metabolized to ethanol, by means of the bifunctional acetaldehyde-CoA/alcohol dehydrogenase (Adh2, EC 1.1.1.1), and/ or to acetate by the activity of the phosphate acetyltransferase (Pta, EC 2.3.1.8). The theoretical ratio of the bifido shunt gives 1 mol of lactic acid and 1.5 moles of acetic acid per mol of glucose consumed, producing 2.5 moles of ATP without CO2 formation, although the production of formic acid and ethanol can modify this fermentation balance. The energetic balance of the bifido shunt (2.5 moles ATP) is higher than that obtained by lactic acid bacteria after the hexose metabolism through the EMP (or homofermentative pathway, 2  moles ATP) or through the pentose-phosphoketolase (or heterofermentative pathway, 1 mol ATP). The availability of several genome sequences of Bifidobacterium species provides genetic evidence to indicate that it is a prototrophic genus for some growth factors—that is, it is able to synthesize some vitamins (B complex, folic acid, thiamine, and nicotinate), aminoacids, and nucleotides. Genomic information also indicates that several species harbor genes required for metabolizing many monosaccharides or disaccharides through the fructose-6-phosphate phosphoketolase pathway. Analysis of the first bifidobacterial genome available, B. longum NCC2705,30 indicated that this strain possesses all the enzyme-coding genes necessary to metabolize several plant- and milk-derived oligosaccharides, containing fructose, galactose, N-acetyl-glucosamine, N-acetyl-galactosamine, arabinose, xylose, ribose, sucrose, lactose, cellobiose, and melibiose through this shunt. In fact, nearly 10% of the genes contained in the genome of this strain encode for proteins that are predicted to be involved in the carbohydrate metabolism and transport. This supposes,

48

Molecular Detection of Human Bacterial Pathogens

Glucose ATP

GlkA Glucose-6-P Gpi Fructose-6-P

Acetyl-P

Xfp Fructose-6-P

Erythrose-1-P

Sedoheptulose-7-P

ATP

Glyceraldehyde-3-P Tkt

Ribose-5-P

AckA

Tal

Xylulose-5-P

Xfp Acetate

Glyceraldehyde-3-P NADH + H+

Gap 1.3-Biphosphoglycerate Pgk

ATP

3-Phosphoglycerate

AckA

Gpm

ATP

2-Phosphoglycerate Eno

Acetyl-P

Phosphoenolpyruvate ATP

Pyk Pyruvate

Pfl

Ldh2

NAD*

Lactate

Pta Acetyl-CoA Adh2

Formate

Acetaldehyde Adh2

EMP pathway

NAD*

NAD*

Ethanol

FIGURE 5.2  Fructose-6-phosphate phosphoketolase pathway “Bifid shunt.”

theoretically, a selective advantage to colonize the colon, an environment poor in mono- and disaccharides because these are consumed by the host and microbiota in the upper GIT. Based on amino acid sequence identity, the presence of several glycolytic activities, including xylanases, arabinosidades, α-galactosidases, neopullanase, isomaltase, maltase, inulinase, β-galactosidases, β-glucosidases, hexosaminidases, and α-manosidases, as well as eight high-affinity oligosaccharide transporters, has been predicted.30 Similar gene arrangements have been seen after the genome annotation and comparison of the bifidobacterial species B. adolescentis ATCC15703, B. animalis subsp. lactis AD011, and B. longum subsp. infantis ATCC15697.31,32 However, comparative genome analysis revealed some differences, such as the presence of a fos gene cluster in B. animalis subsp. lactis AD011, very similar to the one described in B. breve UCC2003.33 This cluster is likely to be involved in fructooligosaccharide (FOS) catabolism, which is present in a panoply of food ingredients and supplements. FOS are composed of 4–60 fructose monomers linked by β(2–1) bonds, with a terminal glucose also connected by β(2–1)

linkage. This β(2–1) bond is not recognized as a substrate by intestinal enzymes, and thus FOS are able to reach the colon, where they stimulate the growth of beneficial bacteria, including bifidobacteria.34 This capacity of beneficially modulating the composition of the intestinal microbiota allows FOS to be considered as prebiotics.35 The presence of fos cluster reflects the metabolic adaptation of bifidobacteria to the human GIT conditions. Interestingly, comparative genomics revealed the presence of a specific genes cluster involved in the transport and metabolism of human milk oligosaccharides (HMO) in the strain B. longum subsp. infantis ATCC15697.31 This strain lacked certain enzymatic activities related to the metabolism of plant-derived oligosaccharides, such as arabinose and xylose, which are abundant in adult diets. This fact, together with the evidence of the presence of similar HMO-related gene clusters in infant gut metagenomes, suggests a specific adaptation of B. longum subsp. infantis ATCC15697 to the infant GIT.36 Thus, bifidobacteria are able to ferment a wide variety of carbohydrates. In addition, they are capable of

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Bifidobacterium

TABLE 5.1 Accession Numbers of the Major Proteins Acting in Bifid Shunt in Some Available Bifidobacterium Genomes

GlkA Gpi Tal Tkt Xfp Gap Pgk Gpm Eno Pyk Ldh2 Pfl Pta AckA Adh2

B. adolescentis ATCC 15703

B. animalis subsp. lactis AD011

B. bifidum NCIMB 41171

B. longum subsp. longum NCC2705

B. longum subsp. infantis ATCC 15697

YP_909820 YP_909094 YP_909693 YP_909692 YP_909550 YP_909942 YP_909698 YP_909241 YP_909508 YP_909541 YP_909980 YP_909855 YP_909551 YP_909552 YP_909182

YP_002470082 YP_002469157 YP_002470219 YP_002470218 YP_002470341 YP_002469609 YP_002470227 YP_002469301 YP_002469894 YP_002470352 YP_002469660 YP_002469833 YP_002470342 YP_002470343 YP_002469236

ZP_03647167 ZP_03645763 ZP_03646448 ZP_03646447 ZP_03646196 ZP_03645995 ZP_03646458 ZP_03647102 ZP_03646131 ZP_03646166 ZP_03646455 ZP_03646658 ZP_03646187 ZP_03646186 ZP_03647027

NP_696841 NP_695484 NP_695898 NP_695899 NP_696135 NP_696527 NP_695890 NP_696806 NP_696193 NP_696160 NP_696472 NP_696127 NP_696142 NP_696143 NP_696730

YP_002322055 YP_002321916 YP_002322563 YP_002322564 YP_002323176 YP_002322378 YP_002322555 YP_002323591 YP_002323289 YP_002323198 YP_002322558 YP_002323169 YP_002323183 YP_002323184 YP_002323678

Additional information can be obtained through their corresponding accession number at http://www.ncbi.nlm.nih.gov/protein. GlkA, glucokinase; Gpi, glucose-6-phosphate isomerase; Tal, transaldolase; Tkt, transketolase; Xfp, xylulose-5-phosphate/fructose-6-phosphate phosphoketolase; Gap, glyceraldehyde-3-phosphate dehydrogenase C; Pgk, phosphoglycerate kinase; Gpm, phosphoglycerate mutase; Eno, enolase; Pyk, pyruvate kinase; Ldh2, lactate dehydrogenase; Pfl, formate acetyltransferase; Pta, Phosphate acetyltransferase; AckA, acetate kinase; Adh2, bifunctional acetaldehyde-CoA/alcohol dehydrogenase.

processing indigestible complex carbohydrates from our diet and from host origin.6,30,37–39 Hence, the availability of genome sequences of intestinal Bifidobacterium spp., together with the use of physiological experiments, has provided considerable insight into the adaptation of this genus to the human gut environment, ­especially regarding the utilization of dietary carbohydrates, resistance to bile and acid conditions, and interaction with the host.

5.1.3  Clinical Features and Pathogenesis It has been shown that Bifidobacterium is present at high levels in the gut microbiota mainly in infants but also in adults.40 Reduced levels of these microorganisms in the intestine of subjects suffering from different diseases, including colon cancer, irritable bowel syndrome, inflammatory bowel disease, atopic disease, or celiac disease, have been observed.41–45 For these reasons several health-promoting properties have been attributed to the microorganisms of this genus and increasing bifidobacterial levels in the GIT is often a treatment target of dietary intervention strategies. So far, there have been no reports of sepsis related to bifidobacteria use in otherwise healthy individuals. Therefore, the current evidence strongly suggests that bifidobacteria are extremely safe to use in the general population. In fact, the most commonly used Bifidobacterium species are considered to be generally safe (GRAS), and in Europe such microorganisms have been granted a Qualified Presumption of Safety (QPS) status.46 Despite the widespread use of the

genus Bifidobacterium, in particular the species B. animalis subsp. lactis, in commercial probiotic products, these probiotics have never been related to sepsis associated with probiotic use, demonstrating the low pathogenic potential of bifidobacteria. This extensive use of bifidobacteria as probiotic agents, together with the numerous clinical trials conducted with these microorganisms, constitutes good proof of safety. As of July 2009, 43 undergoing or completed human clinical trials in which bifidobacteria were administered can be found registered at ClinicalTrials.gov. During last year alone, more than 25 human intervention studies, in which around 2000 volunteers—including healthy adults and elderly, but also full-term and preterm infants, HIV infected children, allergic subjects, pregnant women or IBS patients among others— received a test product containing bifidobacteria, have been published without reporting any detrimental effects. In vitro assessments offer means to investigate the safety of bifidobacteria based on the intrinsic properties of the strains. Several different in vitro approaches have been used in the safety assessment of bifidobacteria. In vitro tests assessing the resistance to antibiotics and the presence of mobile antibiotic resistance genes are common and constitute perhaps the major safety concern regarding bifidobacteria. Several studies have attempted to identify the presence of relevant virulence determinants in bifidobacteria. However, to date such determinants have not been identified. It should be noted, however, that classical risk assessments, which are used for pathogens, might not always be directly applicable to microorganisms such as bifidobacteria, which become members of

50

the normal healthy intestinal microbiota soon after birth and are also components of normal human diet. In pathogens, pathogenicity is normally a consequence of several properties of the strain. The possible presence of such a property in a strain of low infective potential and low clinical significance does not imply that the strain is pathogenic or poses a risk to health.47 An example of this is the ability to adhere to human mucosa, which is a virulence factor in the case of true pathogens, but is also an essential feature of many commensal microbes with very low pathogenic potential, such as bifidobacteria. To date, no clear virulence factors similar to those associated with pathogenic microorganisms have been identified for bifidobacteria.48 This is likely to result from the minimal infectivity of the bifidobacteria. Animal models have been also used in the safety ­evaluation of Bifidobacterium.49,50 Acute oral toxicity tests have been carried out without observing toxicity.49,51 A number of ­studies have examined the effects of bifidobacteria in ­experimentally induced colitis in murine models. Despite disruption of the intestinal barrier in colitic animals, side effects due to increased bacterial translocation seem not to be apparent. In contrast, improvement of clinical condition by reducing the inflammatory colitis of these animals has been often reported. Immunocompromised animal models of both adult and young animals have also been tested to evaluate safety without observing detrimental effects. However, many of the animal models applied in the safety assessment of probiotics were originally developed to study pathogenic microorganisms in which virulence traits are present, and therefore they may not be optimal for studying translocation of non-virulent microorganisms such as bifidobacteria. With regard to antibiotic resistance, Bifidobacterium ­species are resistant to aminoglycosides, metronidazole, and gram-negative spectrum antibiotics. They are also intrinsically resistant to mupirocin. In contrast, bifidobacteria are very susceptible to macrolides/lincosamides, vancomycin, rifampicin, spectinomycin, chloramphenicol, and β-lactams. The susceptibility to tetracyclines and cephalosporins varies among strains.52 One of the current major concerns for the safe use of Bifidobacterium strains is the presence of tetracycline resistance genes, especially tet(W), although other genes, such as tet(M), tet(O), tet(L), tet(W/32/O), and tet(O/W), have been detected, albeit much less frequently.53 In bifidobacteria, tet genes seem to be integrated in the chromosome, and they have not been associated with transposons or plasmids, but they are very often flanked by putative transposase genes.53–55 Transposases are enzymes that catalyze the movement of DNA segments among different locations by recognizing insertion sequences in the DNA, and they are thought to be involved in the mobilization of tet(W) genes in bifidobacteria.54 The long history of safe consumption and the lack of toxicity and virulence determinants support the safety of Bifidobacterium. On the other hand, the low incidence rate of infections due to these microorganisms seems to support this hypothesis. However, it is important to underline that there is no “zero risk” in microbiology and safety assessment is always essential. In fact, although very rare,

Molecular Detection of Human Bacterial Pathogens

some bifidobacteria, probably of intestinal origin, have been ­associated with a few cases of bacteremia, usually in patients with underlying diseases.56 Reduced immune function is the main common denominator in these patients and such patients are prone to bacteremia in general and not ­bifidobacterial ­infection in particular. Of the about 2000 culture-positive cases of cervicofacial actinomycoses assessed by Pulverer et al., none was due to bifidobacteria.57 The few cases of infection or sepsis reported have been mainly related to Bifidobacterium eriksonii (also known as Actinomyces eriksonii) which has been considered a synonym of B. dentium.58 Often these cases are related with subjects with subjacent problems and correspond to ­polymicrobial infections.59 A fatal case of pulmonary infection due to B. eriksonii in a 52-year-old alcoholic patient with ­periodontal disease was reported.60 The presence of bifidobacteria was also observed in a case of infected cavitating lung tumor61 and a case of pericarditis.62 B. adolescentis was isolated in a case of urinary tract infection in a child.63 The presence of B. dentium (named as A. eriksonii), often together with other microorganisms, in some rare cases of abscesses has also been observed.64,65 B. breve was reported to be the causative agent of a neonatal meningitis case, being the second case of bifidobacterial meningitis reported to date.66 Ha et al.67 published a rare case of B. longum septicemia. The patient had undergone acupuncture for a lumbar hernia and small needles had been inserted in the lumbar area and left there. Other than the herniated intervertebral disk the subject was healthy, thus it seems that needle contamination or intestinal punctures by a needle were the likely causes of the infection. Despite these few and isolated cases, the pathogenicity of bifidobacteria appears to be very low. Bourne et al.68 carried out over 90,000 blood cultures from subjects with bacteremia symptoms, recovering over 9000 bacterial isolates. Only 10 out of those isolates were bifidobacteria, and in five cases it was possible to identify the strain as B. eriksonii. All these isolates were obtained from nine patients (in two of the cases, other microorganisms were also found) with underlying conditions (pregnancy complications, gastrointestinal disorders, systemic lupus). To date, the major concern regards caries and mainly B. dentium and Bifidobacterium-related genera such as Parascardovia and Scardovia, as these microorganisms may be found forming part of the complex microbial community present in caries.69,70 Clinical trials using bifidobacteria, along with the ­widespread and safe use of this microbial genus worldwide, constitute the most compelling evidence of the safety of probiotics. Although very often the main target outcome in these intervention trials focuses on the health benefits of the strain, the safety of the administration and the potential adverse events, if any, are often reported as a secondary outcome. Clinical safety trials enable the in vivo evaluation of the effects in humans in a controlled manner, with a special focus on the attributes relevant to the safety of the administered strain and the factors contributing to possible adverse

51

Bifidobacterium

events. However, these clinical investigations are most commonly carried out with healthy volunteers. Considering the infection cases observed, it is clear that safety of bifidobacteria may be of particular interest to specific groups, such as neonates, who have compromised immune systems, or diseased patients. In neonates and low-birth weight infants, successful clinical interventions have been carried out,71 and clinical trials suggest that these microorganisms are safe to use in follow-on formulas and growing-up milks.72 Clinical evaluation in elderly populations is also of special interest, since elderly subjects commonly have health-related problems including infections and gastrointestinal problems. The safety and the lack of adverse events following the consumption of Bifidobacterium strains by elderly subjects have also been demonstrated. However, the possible risks may be elevated in certain high-risk populations, and the demonstration of the safety of a certain strain in the general population does not necessarily imply that the administration of the same strain is equally safe in high-risk populations. In fact, despite the overwhelming evidence supporting bifidobacterial safety, there may be potentially adverse events, as demonstrated by a recent study carried out in severely ill patients with acute pancreatitis. Besselink et al.73 investigated the effects of a probiotic mix containing Lactobacillus, Lactococcus, and Bifidobacterium strains in a randomized placebo-controlled trial that included 296 adults with a first episode of high-risk acute pancreatitis. The probiotic mix was administered by nasojejunal feeding tube, and had been specifically designed, on the basis of previous studies, to inhibit the growth of pathogens important in pancreatic necrosis.74 The authors found a 2.53-fold increase in mortality risk in probiotic-treated participants, and nine cases of bowel ischemia (eight fatal) in the probiotic group, with no bowel ischemia in the placebo group. It is known that small bowel ischemia, increased intestinal permeability, and increased bacterial translocation are all associated with acute pancreatitis. In these cases, the direct application of probiotic bacteria to an already damaged small intestinal mucosa may have precipitated a local inflammatory response leading to increased risk of small bowel ischemia. However, it is not known why exactly mortality was higher in the treatment group, and although the mortality was associated with randomization, this does not necessarily imply that the probiotic itself was the causative factor.75 In these cases, the gastrointestinal system plays a central role in the multisystem organ failure, leading to a breakdown of the intestinal barrier that increases translocation of bacteria and their components into the systemic circulation. In spite of that, translocation is difficult to induce in healthy humans, and if this occurs, detrimental effects have not been demonstrated for bifidobacteria. Due to the general consideration as safe of bifidobacteria, most cases of the presence of these microorganisms in infection sites have been considered as contaminants and, much less probably, as pathogens.76 In any case, despite the excellent overall safety record of bifidobacteria, they should be used with caution in certain specific patient groups with severe underlying diseases and reduced immune function. It

is important to note that such patients in general have a high risk for bacteremia and, very often, intestinal bacteria are the origin of the infection.

5.1.4  Diagnosis Other than the infection cases reported, bifidobacteria may have a profound effect on human health. Low levels of intestinal bifidobacteria population and the misbalance of intestinal microbiota are thought to play an important role in the etiology and pathogenesis of different inflammatory, metabolic, and autoimmune diseases. Thus, alterations in intestinal microbiota composition are found in patients with allergies or those suffering from different health disorders.42,43,77,78 Whether these alterations on the microbiota composition and levels are the cause or consequence of illness is currently far from known. In any case, therapy with bifidobacteria and other probiotics have been successfully used for the alleviation of symptoms of these chronic disorders.79,80 The gut, particularly the colon, is largely the organ containing the highest population and diversity of Bifido­ bacterium species. Owing to this fact, methods for detection and enumeration of these microorganisms have mainly been developed focusing on intestinal bifidobacteria, taking into account the difficulty of isolation and characterization inherent in the high number and diversity of other microorganisms sharing the same environment. These methods have also been subsequently adapted and applied to the study of bifidobacteria in other locations. In this respect, it is worth ­mentioning the recent studies on Bifidobacterium species present in breast milk and the possible transmission of bifidobacterial populations, or their DNA, from mothers to children through ­placenta and breastfeeding.21,81,82 As deduced from that indicated above, although bifidobacteria have not been directly considered as the causative agent of any pathology, the development of quantitative and qualitative methods of detection and identification of these microorganisms has, and will, greatly contribute to gain insight into the relationship between levels and species composition of bifidobacteria populations and the health status of the host. The correct identification and typification of Bifido­ bacterium strains is the starting point to establish its microbiological properties and safety. Thus, it is necessary to use adequate tools to provide proper strain identification, to track Bifidobacterium strains during food production, during their GIT passage and establishment, or to identify potential clinical isolates. 5.1.4.1  Phenotypic Techniques The traditional identification of bifidobacteria has been based on phenotypic features. Cell morphology, production of metabolites, enzymatic activities and sugar fermentation profiles are the most commonly analyzed phenotypic characteristics. The association of a typical branched shape with the presence of fructose-6-phosphate phosphoketolase (F6PPK) activity is the first indication that a strain belongs to the genus Bifidobacterium.83 However, not all strains have

52

a V or clamped shape, and the levels of F6PPK activity vary enormously among species and strains.84 Furthermore, the identification at species/strain level have some additional problems, and the classical phenotyping, such as sugar fermentation profiles, transaldolase serotyping, cell wall composition, and the study of the F6PPK isoforms, is clearly not discriminative enough to reach species, subspecies, and biotype-level identification with accuracy. In fact, most cases of bifidobacteria misidentifications are due to inappropriate phenotypic methods.85 Several different quantitative and qualitative methodologies have been used for Bifidobacterium microbiota assessment.86,87 Traditional methods for the study of qualitative and quantitative composition of intestinal bifidobacteria have been carried out by the cultivation of feces. In some cases, it has also been considered that there are different mucosa-associated intestinal bacteria that can differ from feces in biopsies or samples taken from healthy individuals88 or patients.43,89 Due to the anaerobic nature of bifidobacteria, the samples must be handled in anaerobic conditions and processed immediately or shortly after collection. Bifidobacteria are nutritionally demanding microorganisms and therefore are cultured on suitable complex growth media (such as MRS, Rogosa, Beerens, and trypticase phytone yeast extract). Both nonselective and selective differential media have been used for growth and counting, with and without the addition of selective agents.90–92 The choice of one medium or another is dependent on the other bacteria present in the sample and on the method used for subsequent identification. Besides the characteristic “bifid” cell morphology and the F6PPK activity of the genus Bifidobacterium, another phenotypic method for identification at the species level is based on the determination of the cellular fatty acid composition93 with the help of the chromatographic MIDI system directly in feces42 or in previously isolated cultures.92 In spite of the accuracy of the method, it is important to mention that the composition of fatty acids changes notably in bifidobacteria stressed cells,94 which can lead to misidentifications if other alternative methods were not used in those cases. The difficulty of cultivation of anaerobic intestinal microorganisms meant that until a few years ago, their study was restricted to the cultivable species and among them, only to the viable population. This led to the overestimation of some species and the underestimation of others. In addition, not all Bifidobacterium species are recovered from selective media with the same efficiency,90,91 which could draw a distorted picture of the bifidobacteria population inhabiting complex human ecosystems. 5.1.4.2  Molecular Techniques In the last few years, new molecular tools have been developed and applied, as alternative or complementary methods to traditional cultures, for the identification and typing of bifidobacteria. Most of these techniques use 16S rRNA and its encoding genes as target molecules, on which specific PCR primers and hybridization probes are designed to detect particular species or groups of microorganisms, including

Molecular Detection of Human Bacterial Pathogens

the genus Bifidobacterium95–98 and some of its species.40,96,97 Moreover, the use of 16S rRNA instead of the DNA (by obtaining cDNA from mRNA through reverse-transcriptase PCR reactions) can provide data on the activity and viability of microorganisms in their ecosystem rather than on cellular levels as occurs when DNA is the target molecule. Among the most extensively used molecular tools for qualitative assessment of bifidobacteria is the sequence analysis of amplified 16S RNA genes, by PCR amplification from biological samples, followed by cloning and sequencing of the amplified DNA. Other widely used techniques are TGGE and DGGE, using primers for PCR amplification with one of these primers being tagged to a GC clamp to avoid complete dissociation of DNA strands. Amplified fragments are subsequently separated by denaturing gel electrophoresis through a ­gradient of ­temperature (TGGE) or denaturant agent (DGGE).99 Furthermore, PCR amplification products can be separated by high-performance liquid chromatography (HPLC).100 Analysis of terminal-restriction fragment length polymorphism (T-RFLP) by digestion of fluorescent labeled fragments with suitable restriction enzymes101 or the use of Oligonucleotide arrays102 may be also utilized. Sequencing of ribosomal rRNA genes (rDNA) is nowadays the standard methodology for bifidobacteria identification. The bacterial rRNA genes are organized in rrn operons containing the genes for the three ribonucleic acids (5S RNA, 16S RNA, and 23S RNA), with bifidobacteria harboring from two to six operons, depending on the species.10,30 The 16S and 23S rRNA genes, as well as the 16S23S spacer region (Intergenic Transcribed Sequence, ITS), showed nucleotide fingerprints with different discriminatory levels. Sequencing of the 16S rRNA can be used for identification purposes or to establish phylogenetic relationships among species, subspecies, and biotypes,5,97,103 whereas the 16S-23S rRNA ITS sequence is much more variable than the 16S rRNA structural gene, even within closely related taxa, which makes it a valuable target for species identification by using species-specific primers,104 but also its higher discriminatory capacity may allow the differentiation of strains.105 Recently, multilocus sequence typing (MLST) methods have been applied for the typification of Bifidobacterium species and strains. MLST methods made use of a DNA sequencing procedure to characterize different housekeeping  gene loci in the bacterial genome.106 Its application in bifidobacteria has been shown to be highly discriminatory, the target genes studied for detailed identification and classification purposes down to subspecies level being numerous.5,107–110 Application of a variety of PCR protocols has enabled the differentiation between strains of the same species. Random amplification of polymorphic DNA (RAPD) techniques, amplified ribosomal RNA restriction analysis (ARDRA), ERIC (Enterobacterial Repetitive Intergenic Consensus sequence)-PCR, and REP (Repetitive Extrogenic Palindromic)-PCR are the most common PCR-based methods for bifidobacterial typing.4,14,111–113 Other PCR-based

53

Bifidobacterium

typing approaches include T/DGGE (temperature/denaturant gradient gel electrophoresis), amplified fragment length polymorphism (AFLP), PCR coupled to enzyme-linked immunosorbent assay (ELISA), triplicate arbitrarily primer (TAP)-PCR, restriction fragment length polymorphism (RFLP)-PCR, and multiplex PCR.108,111,114–116 Some other methods for bifidobacteria identification and typing used DNA profiling, such as plasmid analysis, RFLP, or PFGE (pulsed field gel electrophoresis) of total DNA.4,111,117 PFGE is often considered the best technique for strain-­specific typification and has shown a high discriminatory power. Among the nucleic acids hybridization methods, ribotyping, microplate blot, and microarray analysis have also been used for bifidobacterial typing.102,111,118 Finally, it is worthwhile to point out that other methods based on metabolite ­production, have also been applied to the identification and typing of Bifidobacterium such as chromatographic analysis of organic acids or the analysis of the intrinsic fluorescence of aromatic amino acids.119,120 Qualitative techniques provide a picture of the most abundant species and the diversity in microbial ecosystems but fail to provide data on the relative abundance of microbial groups as well as on the less abundant species. Among the quantitative techniques most extensively used for Bifidobacterium is fluorescent in situ hybridization (FISH), with specific probes coupled to flow cytometry or fluorescence microscopy detection. Although this technique has widely been used to study the evolution of intestinal populations,45,95,121 it is extremely laborious and time consuming, and is influenced by cell permeability or the ribosome content of cells. Quantitative real-time PCR is a faster tool to study complex microbial communities. Different PCR assays have been developed for fecal bifidobacteria and its species, or focused on the metabolically active microorganisms targeting 16S rDNA or rRNA, respectively, by using probes labeled with different fluorescent dyes. “Omics” (metagenomics, metaproteomics, and metabolomics) are very recent, high throughput, and promising techniques that can also be applied to the study of bifidobacteria in complex ecosystems. The sensitivity level of genetic methods for identification and typing of bacterial isolates is obviously limited by the previous sensitivity of the culture media and conditions used for isolating the microorganisms.

5.2  METHODS 5.2.1  Sample Preparation Most of the studies evaluating bifidobacterial population ­composition and levels have used fecal samples as starting material. However, the same methods with slight modifications can be used to determine the presence or levels of ­bifidobacteria in other matrixes such as human tissues or fluids. Both culture-dependent and culture-independent methods are available to assess Bifidobacterium populations. Depending on the method used for the detection of the microorganism, different sample preparation procedures are needed.

When a culture-dependent method is going to be used, isolation of the microorganisms is required prior to identification by phenotypic or genotypic techniques. Given the complex microbial community present in fecal samples, isolation of bifidobacteria is a challenging issue. Typically, the fecal sample is diluted in an isotonic buffer (e.g., PBS) and homogenized using a blender. Decimal dilutions are then prepared and 0.1 mL of the appropriate dilution is plated in an adequate selective or differential culture media.91,92 After the incubation of the plates at 37°C for 48 h under anaerobic conditions in an anaerobic chamber or a jar with an anaerobiosis system (e.g., Anaerocult A, Merck), putative bifidobacterial colonies may be picked up and their identity further confirmed by phenotypic or molecular techniques as stated in Section 5.1.4. Very often, molecular methods of identification focus on DNA analyses that require a step of extraction of DNA from the studied strain. Bifidobacteria are not easily lysed, and for this reason lysis buffers are often supplemented with lysozyme and mutanolysin to improve the lysis efficiency, thereby increasing the yield of the DNA extraction. Nowadays, in addition to the traditional methods of DNA extraction using phenol/chloroform, a number of DNA extraction kits are commercially available from the main molecular biology suppliers and are very often used for bacterial DNA extraction. Culture-independent methods do not require a previous bacterial isolation step. In this case, the starting material may be a complex sample (i.e., fecal sample). The total DNA in the sample is extracted previously to the specific detection of bifidobacterial DNA in the complex DNA pool extract. Different DNA extraction methods for such complex samples have been reported in the literature19,99 and commercial kits are also available (e.g., QIAamp DNA Stool Kit, Qiagen) to this end.

5.2.2  Detection Procedures Among the easiest and most currently used molecular tools to detect the presence of bifidobacteria in complex samples is the use of PCR with specific primers. Several specific primers have been designed for the genus Bifidobacterium97,98 and for different bifidobacterial species.40,96,97,122 These primers have been extensively used for the culture-independent detection of bifidobacteria, mainly in fecal samples but also in other matrixes, including food products. Procedure 1. Prepare the PCR mix (most commonly 25 or 50 µL) containing PCR buffer, DNA polymerase, dNTPs, and the appropriate primers (genus or species-specific depending on the aim of the study). Add DNA extract (most often 2–5 µL) or an equal amount of water (negative control). 2. Run the PCR program using the optimal conditions for the DNA polymerase used and the adequate annealing temperature (depending in the primers used).

54



Molecular Detection of Human Bacterial Pathogens

3. After completion of PCR, add DNA loading buffer and visualize amplification products by standard agarose gel electrophoresis and ethidium bromide staining.

Note: Some of these primers have also been used for ­determination of bifidobacterial levels by means of quantitative real-time PCR. Two approaches may be used to this end; a combination of the primers with a DNA dying agent such as SYBR Green (SYBR Green assay),15,21,44 or the combined use of the primers with a labeled internal oligonucleotide probe (5´nuclease assay).40,98,122

5.3  CONCLUSION AND FUTURE PERSPECTIVES Bifidobacteria are normal inhabitants of the human ­intestinal tract, constituting an important part of the microbiota of the colon, where they reach very high numbers. These ­microorganisms have shown a high adaptation to their ­environment and are fastidious and difficult to cultivate in the laboratory. However, the development of techniques to grow anaerobic microorganisms, together with the recent ­development of culture-independent methods, has allowed an improvement of our knowledge on Bifidobacterium ­physiology and ecology. Although some rare cases of infection have been reported, usually in subjects with underlying diseases, the major ­concern about Bifidobacterium safety regards the potential role of B.  dentium in dental caries. In fact, these ­microorganisms have shown a very low pathogenic potential, as has been demonstrated by in vitro animal and human studies. Moreover, several studies have indicated that reduced intestinal bifidobacterial levels may be related to different diseases and ­several ­health-promoting properties have been attributed to these microorganisms. For these reasons, increasing bifidobacterial levels has often been taken as a treatment target. Different nutritional intervention strategies have been developed to this end, mainly using probiotic bacteria, including bifidobacteria, and prebiotic compounds. In this regard, it is important to underline that there is no “zero risk” and that the addition of any microorganisms to products for human or ­animal ­consumption requires a rigorous assessment of its safety. In the near future, the recent developments on ­molecular methods and the availability of bifidobacterial genomes will improve our understanding of the biology of the genus Bifidobacterium, the interaction with the host, and the role in health and disease.





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3. Tissier, M.H., Réchérches sur la flore intestinale normale et pathologique du nourisson (thesis), University of Paris, France, 1900. 4. Satokari, R.M. et al., Molecular approaches for the detection and identification of bifidobacteria and lactobacilli in the human gastrointestinal tract, Syst. Appl. Microbiol., 26, 572, 2003. 5. Ventura, M. et al., Analysis of bifidobacterial evolution using a multilocus approach, Int. J. Syst. Evol. Microbiol., 56, 2783, 2006. 6. Ventura, M. et al., Genomics of Actinobacteria: tracing the evolutionary history of an ancient phylum, Microbiol. Mol. Biol. Rev., 71, 495, 2007. 7. Okamoto, M. et al., Bifidobacterium tsurumiense sp. nov., from hamster dental plaque, Int. J. Syst. Evol. Microbiol., 58, 144, 2008. 8. Killer, J. et al., Bifidobacterium bombi sp. nov., from the ­bumblebee digestive tract, Int. J. Syst. Evol. Microbiol., 59, 2020, 2009. 9. Watanabe, K. et al., Bifidobacterium mongoliense sp. nov., from airag, a traditional fermented mare’s milk product from Mongolia, Int. J. Syst. Evol. Microbiol., 59, 1535, 2009. 10. Ventura, M. et al., Insights into the taxonomy, genetics and physiology of bifidobacteria, Antonie Van Leeuwenhoek, 86, 205, 2004. 11. Felis, G.E., and Dellaglio, F., Taxonomy of lactobacilli and bifidobacteria, Curr. Issues Intest. Microbiol., 8, 44, 2007. 12. Masco, L. et al., Culture-dependent and culture-independent qualitative analysis of probiotic products claimed to contain bifidobacteria, Int. J. Food Microbiol., 102, 221, 2005. 13. Reuter, G., The Lactobacillus and Bifidobacterium microflora of the human intestine: composition and succession, Curr. Issues Intest. Microbiol., 2, 43, 2001. 14. Turroni, F. et al., Exploring the diversity of the bifidobacterial population in the human intestinal tract, Appl. Environ. Microbiol., 75, 1534, 2009. 15. Turroni, F. et al., Microbiomic analysis of the bifidobacterial population in the human distal gut, ISME J., 3, 745, 2009. 16. Guarner, F., and Malagelada, J-R., Gut flora in health and disease, Lancet, 381, 512, 2003. 17. Bäckhed, F. et al., Host-bacterial mutualism in the human intestine, Science, 307, 1915, 2005. 18. Isolauri, E., Salminen, S., and Ouwehand, A.C., Probiotics, Best Pract. Res. Clin. Gastroenterol., 18, 299, 2004. 19. Eckburg, P.B. et al., Diversity of the human intestinal microbial flora, Science, 308, 1635, 2005. 20. Beighton, D. et al., Isolation and identification of Bifido­ bacteriaceae from human saliva, Appl. Environ. Microbiol., 74, 6457, 2008. 21. Martin, R. et al., Isolation of bifidobacteria from breast milk and assessment of the bifidobacterial population by PCRdenaturing gradient gel electrophoresis and quantitative realtime PCR, Appl. Environ. Microbiol., 75, 965, 2009. 22. Lamendella, R. et al., Bifidobacteria in feces and environmental waters, Appl. Environ. Microbiol., 74, 575, 2008. 23. Masco, L. et al., In vitro assessment of the gastrointestinal transit tolerance of taxonomic reference strains from human origin and probiotic product isolates of Bifidobacterium, J. Dairy Sci., 90, 3572, 2007. 24. Cotter, P.D., and Hill, C., Surviving the acid test: Responses of gram-positive bacteria to low pH, Microbiol. Mol. Biol. Rev., 67, 429, 2003. 25. Matsumoto, M., Ohishi, H., and Benno, Y., H+-ATPase activity in Bifidobacterium with special reference to acid tolerance, Int. J. Food Microbiol., 93, 109, 2004.

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56 65. Civen, R., Vaisanen, M.L., and Finegold, S.M., Peritonsilar abscess, retropharyngeal adscess, mediastinitis, and nonclostridial anaerobic myonecrosis: A case report, Clin. Infect. Dis., 16, S299, 1993. 66. Nakazawa, T. et al., Neonatal meningitis caused by Bifidobacterium breve, Brain Dev., 18, 160, 1996. 67. Ha, G.Y. et al., Case of sepsis caused by Bifidobacterium longum, J. Clin. Microbiol., 37, 1227, 1999. 68. Bourne, K.A. et al., Bacteremia due to Bifidobacterium, Eubacterium or Lactobacillus; twenty-one cases and review of the literature, Yale J. Biol. Med., 51, 505, 1978. 69. Chhour, K-L. et al., Molecular analysis of microbial diversity in advanced caries, J. Clin. Microbiol., 43, 843, 2005. 70. Mantzourani, M., Fenlon, M., and Beighton, D., Association between bifidobacteriaceae and the clinical severity of root caries lesions, Oral Microbiol. Immunol., 24, 32, 2009. 71. Hoyos, A.B., Reduced incidence of necrotizing ­enterocolitis associated with enteral administration of Lactobacillus acidophilus and Bifidobacterium infantis to neonates in an intensive care unit, Int. J. Infect. Dis., 3, 197, 1999. 72. Haschke, F. et al., Clinical trials prove the safety and efficacy of the probiotic strain Bifidobacterium Bb12 in follow-up ­formula and growing-up milks, Monatsschr. Kinderheilkd., 1146, S26, 1998. 73. Besselink, M.G.H. et al., Probiotic prophylaxis in predicted severe acute pancreatitis: A randomized, double-blind, placebocontrolled trial, Lancet, 371, 651, 2008. 74. van Minnen, L.P. et al., Modification of intestinal flora with multispecies probiotics reduces bacterial translocation and improves clinical course in a rat model of acute pancreatitis, Surgery, 141, 470, 2007. 75. Reid, G. et al., Probiotic prophylaxis in predicted severe acute pancreatitis, Lancet, 372, 112, 2008. 76. Liong, M.T., Safety of probiotics: translocation and infection, Nutr. Rev., 66, 192, 2008. 77. Sanz, Y. et al., Differences in faecal bacterial communities in coeliac and healthy children as detected by PCR and denaturing gradient gel electrophoresis, FEMS Immunol. Med. Microbiol., 51, 562, 2007. 78. Swidsinski, A. et al., Active Crohn’s disease and ulcerative colitis can be specifically diagnosed and monitored based on the biostructure of the fecal flora. Inflamm. Bowel Dis., 14, 147, 2008. 79. Picard, C. et al., Review article: Bifidobacteria ans probiotic agents-physiological effects and clinical benefits, Aliment. Pharmacol. Ther., 22, 495, 2005. 80. Zuccotti, G.V. et al., Probiotics in clinical practice: An ­overview, J. Int. Med. Res., 36, 1A, 2008. 81. Gueimonde, M. et al., Breast milk: A source of bifidobacteria for infant gut development and maturation? Neonatology, 92, 64, 2007. 82. Satokari, R. et al., Bifidobacterium and Lactobacillus DNA in the human placenta, Lett. Appl. Microbiol., 48, 8, 2009. 83. Ballongue, J., Bifidobacteria and probiotic action, In Salminen,  S., von Wright, A., and Ouwehand, A. (Editors), Lactic Acid Bacteria: Microbiological and Functional Aspects, 3rd ed., pp. 67–124S, Marcel Dekker, New York, 2004. 84. Orban, J.I., and Patterson, J.A., Modification of the phosphoketolase assay for rapid identification of bifidobacteria, J. Microbiol. Methods, 40, 221, 2000. 85. Huys, G. et al., Accuracy of species identity of commercial bacterial cultures intended for probiotic or nutritional use, Res. Microbiol., 157, 803, 2006.

Molecular Detection of Human Bacterial Pathogens 86. Gueimonde, M., and de los Reyes-Gavilán, C.G., Detection and enumeration of gastrointestinal microorganisms, In Lee, Y.K., and Salminen, S. (Editors), Handbook of Probiotics and Prebiotics, 2nd ed., pp. 25–43, John Wiley and Sons, Inc., New Jersey, USA, 2009. 87. Margolles, A., Mayo, B., and Ruas-Madiedo, P., Screening, identification, and characterization of Lactobacillus and Bifidobacterium strain. In Lee, Y.K., and Salminen, S. (Editors), Handbook of Probiotics and Prebiotics, 2nd ed., pp.  4–24, John Wiley and Sons, Inc., New Jersey, USA, 2009. 88. Delgado, S., Suárez, A., and Mayo, B., Bifidobacterial diversity determined by culturing and by 16S rDNA sequence analysis in feces and mucosa from ten healthy Spanish adults, Dig. Dis. Sci., 51, 1878, 2006. 89. Conte, M.P. et al., Gut-associated bacterial microbiota in paediatric patients with inflammatory bowel disease, Gut, 55, 1760, 2006. 90. Silvi, S., Rumney, C.J., and Rowland, I.R., An assessment of three selective media for bifidobacteria in faeces, J. Appl. Bacteriol., 81, 561, 1996. 91. Apajalahti, J.H.A. et al., Selective plating underestimates abundance and shows differential recovery of bifidobacterial species from human feces, Appl. Environ. Microbiol., 69, 5731, 2003. 92. Woodmansey, E. et al., Comparison of compositions and ­metabolic activities of fecal microbiotas in young adults and in antibiotic-treated and non-antibiotic treated elderly subjects, Appl. Environ. Microbiol., 70, 6113, 2004. 93. Eerola, E., and Lehtonen, O.P., Optimal data processing procedure for automatic bacterial identification by gas-liquid chromatography of cellular fatty acids. J. Clin. Microbiol., 26, 1745, 1998. 94. Ruiz, L. et al., Cell envelope changes in Bifidobacterium animalis ssp. lactis as a response to bile, FEMS Microbiol. Lett., 274, 316, 2007. 95. Langendijk, P.S. et al., Quantitative fluorescence in-situ hybridization of Bifidobacterium spp. with genus-specific 16S ribosomal-RNA-targeted probes and its application in fecal samples, Appl. Environ. Microbiol., 61, 3069, 1995. 96. Kaufmann, P. et al., Identification and quantification of Bifidobacterium species isolated from food with genus-specific 16S rRNA-targeted probes by colony hybridization and PCR, Appl. Environ. Microbiol., 63, 1268, 1997. 97. Matsuki, T., Watanabe, K., and Tanaka, R., Genus- and ­species-specific PCR primers for the detection and identification of bifidobacteria, Curr. Issues Intest. Microbiol., 4, 61, 2003. 98. Gueimonde, M. et al., New real-time quantitative PCR procedure for quantification of bifidobacteria in human fecal samples, Appl. Environ. Microbiol., 70, 4165, 2004. 99. Favier, C.F., de Vos, W.M., and Akkermans, A.D.L., Development of bacterial and bifidobacterial communities in feces of newborn babies, Anaerobe, 9, 219, 2003. 100. Goldenberg, O. et al., Molecular monitoring of the intestinal flora by denaturing high performance liquid chromatography, J. Microbiol. Meth., 68, 94, 2007. 101. Sakata, S. et al., Culture-independent analysis of fecal microbiota in infants, with special reference to Bifidobacterium species, FEMS Microbiol. Lett., 243, 417, 2005. 102. Boesten, R.J., Schuren, F.H., and de Vos, W.M., A Bifido­ bacterium mixed-species microarray for high resolution discrimination between intestinal bifidobacteria, J. Microbiol. Methods, 76, 269, 2009.

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57 species-specific amplified ribosomal DNA restriction analysis (ARDRA), FEMS Microbiol. Ecol., 36, 113, 2001. 113. Krizova, J., Spanova, A., and Rittich, B., Evaluation of amplified ribosomal DNA restriction analysis (ARDRA) and species-specific PCR for identification of Bifidobacterium species, Syst. Appl. Microbiol., 29, 36, 2006. 114. Cusick, S.M., and O’Sullivan, D.J., Use of a single, triplicate arbitrarily primed-PCR procedure for molecular fingerprinting of lactic acid bacteria, Appl. Environ. Microbiol., 66, 2227, 2000. 115. Favier, C.F. et al., Molecular monitoring of succession of bacterial communities in human neonates, Appl. Environ. Microbiol., 68, 219, 2002. 116. Delcenserie, V. et al., Discrimination between Bifidobacterium species from human and animal origin by PCR-restriction fragment length polymorphism, J. Food Prot., 67, 1284, 2004. 117. Alvarez-Martín, P., Flórez, A.B., and Mayo, B., Screening for plasmids among human bifidobacteria species: Sequencing and analysis of pBC1 from Bifidobacterium catenulatum L48, Plasmid, 57, 165, 2007. 118. Wang, R.F. et al., DNA microarray analysis of predominant human intestinal bacteria in fecal samples, Mol. Cell. Probes, 18, 223, 2004. 119. Lee, K.Y., So, J.S., and Heo, T.R., Thin layer chromatographic determination of organic acids for rapid identification of ­bifidobacteria at genus level, J. Microbiol. Meth., 45, 1, 2001. 120. Ammor, M.S. et al., Reagentless identification of human ­bifidobacteria by intrinsic fluorescence, J. Microbiol. Meth., 69, 100, 2007. 121. Saulnier, D.M.A., Gibson, G.R., and Kolida, S., In vitro effects of selected synbiotics on the human faecal microbiota composition, FEMS Microbiol. Ecol., 66, 516, 2008. 122. Haarman, M., and Knol, J., Quantitative real-time PCR assays to identify and quantify fecal Bifidobacterium species in infants receiving a prebiotic infant formula, Appl. Environ. Microbiol., 71, 2318, 2005.

6 Corynebacterium Luis M. Mateos, Michal Letek, Almudena F. Villadangos, María Fiuza, Efrén Ordoñez, and José A. Gil

CONTENTS 6.1 Introduction........................................................................................................................................................................ 59 6.1.1 Classification, Morphology, and Biology............................................................................................................... 59 6.1.2 Phenotypic and Phylogenetic Characterization...................................................................................................... 60 6.1.2.1 Phenotypic Analysis................................................................................................................................ 60 6.1.2.2 Protein Analysis....................................................................................................................................... 60 6.1.2.3 Phylogenetic Analysis.............................................................................................................................. 64 6.1.3 Pathogenesis and Diagnosis.................................................................................................................................... 65 6.1.3.1 Corynebacterium Diphtheriae................................................................................................................ 65 6.1.3.2 Corynebacterium Species of the Lipophilic Group................................................................................. 66 6.1.3.3 Corynebacterium Species of the Nonlipophilic Group........................................................................... 67 6.2 Methods.............................................................................................................................................................................. 68 6.2.1 Sample Preparation................................................................................................................................................. 68 6.2.1.1 Culture Media for Corynebacterium....................................................................................................... 68 6.2.1.2 DNA Isolation.......................................................................................................................................... 68 6.2.1.3 RNA Isolation.......................................................................................................................................... 69 6.2.2 Detection Procedures.............................................................................................................................................. 69 6.2.2.1 Biolog (GP2) System................................................................................................................................ 69 6.2.2.2 API Coryne System................................................................................................................................. 69 6.2.2.3 Identifications Based on 16S rRNAs Genes............................................................................................ 70 6.3 Future Perspectives and Conclusions................................................................................................................................. 70 Acknowledgments........................................................................................................................................................................ 71 References.................................................................................................................................................................................... 71

6.1  INTRODUCTION 6.1.1  Classification, Morphology, and Biology The coryneform bacteria are a broad group of gram-positive, irregularly shaped rods (sometime oval- or club-shaped), aerobic or facultatively anaerobic, asporogenous, and nonpartially-acid-fast microorganisms classically used in the production of primary metabolites (amino acids, nucleotides, vitamins, etc.).1 The genus Corynebacterium was created in 1896 in order to assign a classification to Corynebacterium diphtheriae (the diphtheroid bacilli), as well as several animal pathogens and parasitic microorganisms.2 During most of the twentieth century, many microorganisms were included in this group based on their staining, morphological, and metabolic properties; in fact, in Bergey’s Manual of Determinative Bacteriology, the coryneform group comprised a broad and diverse number of bacteria of medical

(saprophytic animal and human pathogens), agricultural (plant pathogens), and industrial (saprophytic nonpathogenic amino-acid-producing strains) interest. In the 1990s, phylogenetic (based on 16S and 23S rDNA sequence analyses) and chemotaxonomic (cell wall composition and lipid profiles) analyses resulted in the separation of some members of the coryneform group from those corresponding to plant pathogens.3 Nonetheless, the number of species described for the genus Corynebacterium expanded, from 17 in the first edition of Bergey’s Manual of Systematic Bacteriology4 to more than 70 in the second edition (http://www.bergeys.org/). This large expansion of a bacterial genus is a rare occurrence in microbiology and reflects the enormous interest in these bacteria, mainly in medicine but also by industry. While very few data are available concerning the numbers of corynebacterial species present in nature, given what is known thus far about the genus Corynebacterium, it can be reasonably assumed 59

60

that corynebacteria are abundantly disseminated within a broad range of different natural habitats. In turn, much species diversity remains to be identified and characterized. The most prevalent human pathogen among the corynebacteria is C. diphtheriae, which continues to be a major cause of morbidity and mortality in undeveloped countries and increasingly in developing countries, where nontoxigenic members cause additional pathologies.5 Corynebacterium species other than those belonging to the C. diphtheriae group can be opportunistic or nosocomial pathogens of humans.6 Importantly, infections by these species are becoming increasingly common, mainly due to the remarkable increase in the number of susceptible individuals, especially patients with autoimmune disorders or undergoing immunosuppressive therapy as well as organ-transplanted patients. Species belonging to the genus Corynebacterium constitute a reasonably monophyletic group from the suborder Corynebacterineae, in the order Actinomycetales. They have a GC content of 51%–65% and an optimal temperature for growth of 28°C–37°C. Two broad groups of species from the genus can be distinguished based on their habitats and the consequences of the host-bacteria interaction: (i) Saprophytic members (nonpathogenic) are widely disseminated in different environments, such as soil, sea, dairy products, and animals. Of these, the main species used in industry is C. glutamicum. Additional species include C. ammoniagenes, C.  casei, C. callunae, C. efficiens, C. flavescens, and C. thermoaminogenes.7 (ii) Among the Corynebacterium species of medical importance, the best described is C. diphtheriae, but during the last few decades dozens of others have been recognized as emergent human and animal pathogens. Their optimal temperature for growth is 35°C–37°C and most are auxotrophic, with requirements for vitamins, amino acids, and/or nitrogen bases. Frequently, two broad subgroups of Corynebacterium members are distinguished based on an essential requirement for a particular nutrient, better growth in the presence of lipids (lipophilic group), or equivalent growth in the presence or absence of lipids (nonlipophilic group). Among the lipophilic group, species such as C. accolens, C. afermentans (lipophilum), C. appendicis, C. genitalium, C. hansenii, C. jeikeium, C. kroppenstedtii, C. lipophiloflavum, C. macginleyi, C. pseudogenitalium, C. resistens, C. sputi, C. tuberculostearicum, C. urealyticum, and C. ureicelerivorans are the most representative. Within the nonlipophilic group, a broader number of human pathogens (ascertained or potential) have been described, including C. diphtheriae, C. amycolatum, C. argentoratense, C. atypicum, C. auris, C. aurimucosum, C. confusum, C. coyleae, C. durum, C. freneyi, C. genitalum, C. glucuronolyticum, C. imitans, C. minutissimum, C. mucifaciens, C.  pilbarense, C.  pseudodiphtheriticum,

Molecular Detection of Human Bacterial Pathogens

C.  pseudotuberculosis, C.  resistens, C. riegelii, C.  simulans, C. singulare, C. striatum, C. sundvallense, C. thomssenii, C. timonense, C. tuscanie, C. ulcerans, and C. xerosis. In addition, the nonlipophilic group can be subdivided in fermentative and nonfermentative groups depending, respectively, on the ability or inability (to metabolize sugars under anoxic conditions). Table 6.1 summarizes most of the human-pathogenic Corynebacterium species isolated from clinical samples and characterized thus far, together with the most relevant information concerning these species.

6.1.2  Phenotypic and Phylogenetic Characterization 6.1.2.1  Phenotypic Analysis In-depth characterization of new isolates as members of Corynebacterium and their description as species of this genus rely on phenotypic analyses and thus on a combination of (i) morphological, (ii) physiological, and (iii) chemotaxonomic criteria. Preliminary identification has been traditionally achieved by Gram staining, which reveals the distinctive morphology of Corynebacterium together with additional determination of the bacterium’s physiological properties, such as growth in specific culture media, colony size and shape, pigmentation, cell motility, temperature, and pH. Further analyses include biochemical and enzymatic reactions, such as those determined in automated systems for rapid species identification. The three most frequently used systems are: (i) API Coryne (version 2.0; bioMérieux, France), (ii) BIOLOG (Biolog, Inc., Hayward, CA), and (iii)  RapID CB Plus (Remel, Inc., Norcross, GA). Most of the biochemically based phenotypic analyses are carried out with the test included in the API Coryne system. A brief description of the test’s reactions and corresponding protocol is provided in Section 6.2.2.2, and a description of the protocol for the BIOLOG system in Section 6.2.2.1. The performances of these systems have been evaluated in different studies with variable results because in most cases additional biochemical assays specific for corynebacteria, such as DNase activity and tyrosine hydrolysis, are necessary for definitive identification of the isolates in question.47–49 Most of the chemotaxonomic analyses exploit the presence in corynebacteria of the isomeric form of diaminopimelic acid50 or the bacterial content of whole-cell sugars51 or lipids. Alternatively, they are based on the bacteria’s fatty acids profile or the presence of corynomycolic acids and other β-hydroxy-α-branched carboxylic acids.52 6.1.2.2  Protein Analysis Multilocus Enzyme Electrophoresis (MEE). This technique allows the detection of amino acid substitutions in cellular enzymes that affect their conformation and overall charge. Mobility variants (electromorphs) of the same enzyme can be visualized by their activity in a gel (i.e., as bands of altered migration rates), and each electromorph

Lipo L

L NL NL L

NL NL NL

NL

NL NL NL NL

L NL L NL

Descriptions

<10

<10

>50

<10

<10

<10

>50

<10

<10

<10

>200

<10

<10

<10

<10

<10 <10

C. accolens

C. afermentans (subsp lipophilum)s C. amycolatum

C. argentoratense

C. appendicis

C. atypicum

C.auris

C. aurimucosum

C. confusum

C. coyleae

C. diphtheriae

C. durum

C. freneyi

C. genitalium

C. glucuronolyticum

C. hansenii C. imitans

Corynebacterium Species

+++ ++

++

NF*/+

++

++

++

NF*/+

+

++

NF

+++

NF

+

NF*/+

NF

++

Fermentation

Gi03314 Gi03315

Gi03343/2.8 Md

Gi02702/2.3 Mb

ND

Gi03719

Gc00173/2.8 Mb

Gi05148

Gi03300

Gc00991/2.7 Md

ND

Gi03105

Gi03635

ND

Gi03352/2.5 Md

ND

Gi02699/2.4 Mb

HMP Sequence/Size (Md)

ND ND

59

62

ND

55

53

62/64

ND

60

70

ND

ND

60

58

68

59

GC%

TABLE 6.1 Corynebacterium Species Described as Human Pathogens (or Potential Pathogens)

Type IV CM+ CM+ Tubst– CM+ CM+

Type IV CM+

Type IV CM+ CM– Tubst– Type IV CM+ Type IV CM+ Tubst+ CM– Tubst– Type IV CM+ Type IV CM+ Tubst+ Type IV CM+ Tubst+ Type IV CM+ Type IV CM+ CM+

Type IV CM+

Chemotype

Liposarcom Human throat

Genital tract

Urinary tract/Urethritis

Ulcers/Pus

Respiratory infection

Respiratory infection

Oral/Blood

Foot/Blood

Bronchitis

Ear infection

Clinical sample

Oral infection

Throat

Sepsis/Endocard

Abcess/Valves infect.

Oral tract Endocard

Locus/Disease

Catalase+ Catalase+

Urease– Nit-Red– Catalase+

Catalase+ Nit-Red+ Catalase+

Catalase+

Catalase+

Catalase+ Nit-Red+

Catalase+ Halotolerant Catalase+ Nit-Red– Catalase+

Catalase+ Urease– Catalase+

Catalase+

Catalase+ Urease– Nit-Red+ Urease–

Phenotype

DNA Hyb 16S rDNA DNA Hyb Ribotyping PFGE 16S rDNA

DNA Hyb 16S rDNA 16S rDNA DT-PCR-amp DNA Hyb 16S rDNA DNA Hyb ITS-PCR 16S rDNA DNA Hyb

16S rDNA

DNA Hyb 16S rDNA

16S rDNA

16S rDNA

DNA Hyb 16S rDNA 16S rDNA

ARDRA DNA Hyb 16S rDNA

ARDRA DNA Hyb

Genotype

continued

22 23

21

20

19

18

2

17

16

15

14

13

12

11

10

9

8

Reference

Corynebacterium 61

Lipo L

L L L

NL

NL

NL

NL

NL L NL

L NL NL NL NL

Descriptions

>100

<10

<10

<10

<10

>50

<10

<10

>50

<10

>200

<10

<10

<10

<10

>100

C. jeikeium

C. kroppenstedtii

C. lipophiloflavum

C. macginleyi

C. massiliense

C. minutissimum

C. mucifaciens

C. pilbarense

C. pseudodiphtheriticum

C. pseudogenitalium

C. pseudotuberculosis

C. resistens

C. riegelii

C. simulans

C. singulare

C. striatum

Corynebacterium species

+

++

++

+

+

++

+

NF

+

NF

NF*/+

NF

+

++

++

+

Fermentation

Gi02710/2.7Mb

Gi03319

Gi04645

Gi03318

Gi03107

Gi04675/2.3Mb

Gi02707/2.6Mb

Gi04695

ND

Gi03722

Gi04644

ND

Gi03316

Gi02705/2.3Mb

Gc00770/2.4Mb

Gc00271/2.5 Mb

HMP Sequence/Size (Md)

59

ND

ND

ND

55

52

59

55

ND

64

58/60

ND

58

65

57

61

GC%

TABLE 6.1  (Continued) Corynebacterium Species Described as Human Pathogens (or Potential Pathogens)

Type IV CM+ Type IV+ CM+ Type IV CM+

Tubst+

Type IV CM+ Type IV CM+

Type IV CM+ Tubst– Type IV

Type IV CM+ Tubst+ Type IV CM+ Tubst+ CM+ Tubst+

CM– Tubst+ Type IV CM+ Type IV CM+

CM+

Chemotype

Locus/Disease

Urogenital tract

Semen specimen

Abcess and boil

Urinary infection

Clinical samples

Lynphaden/Granulom

Pulmonary/Oral infection Urinary tract

Clinical ankle aspirate

Oral infection

Clinical sample

Knee arthroplasty

Keratitis corneal/Oral tract

Vaginosis

Sputum

Nosoco-meningitis

Phenotype

Catalase+ Nit-Red+

Catalase+

Catalase+ Urease+ Catalase+

Catalase+ Urease+ Urease– Nit-Red– Catalase+ Oxidase– Urease+ Catalase+

Catalase+ Urease– Nit-Red– Catalase+ Oxidase–

Catalase+

Catalase+ Urease– Catalase+ Urease– Nit-Red+ Catalase+ Oxidase–

Catalase+ Urease– Nit-Red– Catalase+

Genotype

DNAHyb 16S rRNA DNAHyb 16S rRNA 16S rRNA ARDRA DNAHyb 16S rRNA 16S rRNA ARDRA

16S rRNA DT-PCR-amp

16S rRNA

16S rRNA

DNA Hyb 16S rRNA

16S rRNA

DNA Hyb 16S rRNA

16S rRNA rpoB-ampl

16S rRNA

16S rRNA

16S rRNA

DNA Hyb 16S rRNA

38

37

36

35

34

33

32

2

31

30

15,29

28

27

26

25

24

Reference

62 Molecular Detection of Human Bacterial Pathogens

NL NL NL

L

NL NL L

L

NL

<10

<10

<10

<10

<10 >100

>100

<10

>100

C. sundsvallense

C. thomssenii

C. timonense

C. tuberculostearicum

C. tuscanie C. ulcerans

C. urealyticum

C. ureicelerivorans

C. xerosis

+++

+

NF*/+

++ ++

F

++

+++

+++

+

Gi04647

Gi03321

Gi01607/2.3Mb

ND ND

Gi03351/2.4Mb

ND

ND

Gi04426

ND

ND

ND

64

ND ND

60

ND

ND

64

ND

CM+ Type IV CM+ Type IV CM+ Tubst+ Type IV CM+ Tubst+ Type IV CM+

Type IV CM+ Tubst+ Type IV CM+ Type IV CM+ Type IV CM+ Tubst– CM+ Tubst+

Endocard

Septicemia Blood

Cystitis Pyelitis

Blood Endocard Clinical samples

Clinical samples

Endocard

Pleural fluids

Clinical samples

Sputum

Catalase+ Urease– Oxidase– Catalase+ Catalase+ Urease+ Catalase+ Oxidase– Urease+ Catalase+ Urease+ Nit-Red– Catalase+ Urease–

Catalase+ Oxidase– Urease+ Catalase+ Urease+ Catalase+ Urease+ Catalase+ Oxidase–

16S rRNA ARDRA

16S rRNA rpoB-ampl

16S rRNA DNA Hyb

16S rRNA 16S rRNA

PAGE 16S rRNA

16S rRNA rpoB-ampl

PAGE 16S rRNA 16S rRNA

16S rRNA

2

46

45

43 44

42

28

41

40

39

Notes: Descriptions, number of References in PubMed; Lipo, L: lipophilic; NL: Nonlipophilic; fermentation, +++/++/+: relative amount of fermented substrates; NF, nonfermenters or variable*; HMP sequence/ size, code from Human Microbione Project: http://www.hmpdacc.org/; chemotype, type IV: cell wall with meso-diaminopimelic acid, arabinose, and galactose; CM+/–, Corinomycolic acids presence or absence; Tubst+/-, tuberculostearic acids presence or absence; locus-disease, most frequent; phenotype, most characteristic; genotype, molecular technique used for description; reference, number from the references; ND, not determined.

L

<10

C. sputi

Corynebacterium 63

64

potentially represents a distinct allele of that enzyme. The electromorph profile can be used to define the electrophoretic pattern of a corynebacterial strain, assuming that the number of bands analyzed is sufficient to allow intraspecies discrimination. The data are represented as a dendrogram and the distance between the strains determined. This method was used, for example, to distinguish different C. diphtheriae isolates.53 Protein Fingerprinting. SDS-PAGE protein patterns, from whole-cell soluble proteins or a purified particular fraction, can be generated for the strains under study and compared with those of reference strains. Strain-specific differentiation was previously used to characterize certain Corynebacerium strains in clinical isolates. Nowadays, however, it provides supplementary information to that gained through polyphasic approaches in order to fine-tune species identification. 6.1.2.3  Phylogenetic Analysis Over the last two decades, a much broader range of taxonomic studies of bacteria has gradually replaced classical techniques. Molecular analyses of the informational units of cells (proteins, DNA and RNA) have been developed and used in the identification of Corynebacterium members. Modern molecular techniques are based on DNA analysis (DNA–DNA hybridization after gel electrophoresis, the presence of plasmids or integrated elements of uncertain origin, etc.), 16S-23S rRNA gene sequencing (full or partial), PCR-mediated amplification, and protein studies. Different genotyping techniques may be applied to Corynebacterium as tools for either species identification or the differentiation of strains within a species. DNA-based typing methods are highly discriminatory and can be used to distinguish closely related strains with similar phenotypic features. The most frequently used approach is based on PCR-mediated amplification of DNA units common to all bacteria but with internal specific changes. This polymorphism allows the separation of isolates belonging to closely related genera or of members of the same genus but from different species. Identification methods based on 16S rRNA and DNA-dependent RNA polymerase β -subunit (rpoB) genes are mostly employed together with “broad-range” primers.54 However, the analysis of either 16S rRNA or rpoB often lacks specificity due to the absence of polymorphisms. Therefore, many clinical laboratories have sought to design specific primers and the respective protocols to specifically amplify a defined target within one species and thus, consequently, excluding other species.55 In short, limited homology between target genes and broad-range primers on the one hand and insufficient polymorphism among closely related bacteria on the other can preclude the distinction of Corynebacterium members. Perhaps the most complete genotypic analyses of Corynebacterium species to date have been those of Khamis et al.56,57 in which the effectiveness of sequencing the complete 16S rRNA was compared with the use of potentially more discriminatory targets, such as internal fragments of rpoB.

Molecular Detection of Human Bacterial Pathogens

(i) DNA-Based Analysis  Single-Strand Conformation Polymorphism (SSCP). In this classical form of mutation analysis, amplified double-stranded DNA is denatured to generate single-stranded DNA, which is then electrophoresed in a non-denaturing polyacrylamide gel. The migration rate of single-stranded DNA depends not only on its length but also on its secondary structural conformation, resulting in a specific SSCP pattern for each mutant gene. SSCP has been used to analyze important genes of Corynebacterium, such as tox and dtxR.58 Restriction Enzyme Analysis (REA) and Pulse-Field Gel Electrophoresis (PFGE). The digestion of chromosomal DNA (total DNA) with restriction endonucleases generates a series of fragments that are then usually separated by conventional agarose gel electrophoresis. By selecting the appropriate restriction enzyme or a set of enzymes species- or strain-significant, REA patterns are generated. Nonetheless, the resulting banding pattern (fragment sizes between 0.5 and 20 kb) is sufficiently complex such that additional analytic methods are usually required. A modification of REA is pulse-field gel electrophoresis (PFGE), in which a periodic change of the orientation of the electric field enables the separation of high-molecular-weight fragments. PFGE can be combined with the use of rare-cutting restriction endonucleases, which generate a low number of fragments, thus yielding a banding pattern that is more easily interpreted. The typical PFGE DNA fingerprint consists of 5–20 large (10–800 kb), well-resolved fragments and has high discriminatory and reproducible power. Hybridization Analysis. Southern blot assays are used for species identification and are based on the use of randomly cloned heterologous or homologous DNA fragments as probes, which are then hybridized with total DNAs from the new isolates. Most of the newly identified pathogenic Corynebacterium species were initially characterized using other techniques, but hybridization analyses in which the isolates’ total DNA is compared with that of closely related species remains the definitive test to confirm a new species. Ribotyping and Restriction Fragment Length Poly­ morphism (RFLP). Ribotyping is based on the hybridization of digested chromosomal DNA with probes derived from specific rRNA genes. In the genomes of many bacteria, rRNA operons (rrn) are present in several copies and the resulting multiband patterns are frequently strain specific. Although the entire rrn operon can serve as a probe, nowadays a mixture of a few oligonucleotides from conserved regions of 16S–23S rRNA genes is used for this purpose, making the method a truly universal approach to bacterial strain typing. Two genetic patterns are considered identical when they are composed of the same number of fragments and, for homologous fragments, size differences are below a 5% threshold error value. Fingerprint patterns are more stable, more easily reproducible, and more readily interpreted than those obtained with REA. Ribotyping has been used with some success to study the diversity of Corynebacterium strains, although with low discriminatory power. The choice of a suitable restriction enzyme is important and different

65

Corynebacterium

ribopatterns may be obtained by selecting different restriction enzymes. RFLP techniques are equivalent to ribotyping, the difference being that they make use of any DNA sequence (homologous or heterologous) for hybridization purposes. 16S-23S rRNA Gene-sequencing Analyses. All available Corynebacterium 16S rRNA, 23S rRNA, and 16S-23S rRNA spacer region sequences can be retrieved from two databases, EMBL and GenBank, and aligned using the Clustal program. Thus, potentially strain- and species-specific primers can be employed for specific purposes. The amplified rRNA fragment is further sequenced according to conventional techniques and the results are compared with the above-mentioned databases. As this is the molecular technique most widely used in Corynebacterium species identification and determination, a detailed protocol is described in the Methods section of this chapter. Multilocus Sequence Typing (MLST). This method is based on the sequential amplification of several housekeeping genes (e.g., adk, dnaA, fumC, gltA, gyrB, icd, purA) obtained from the whole-genome sequences of four Corynebacterium species (C. glutamicum, C. diphtheriae, C. efficiens, and C. jeikeium). The design of a primer pair for each of the target genes is based on seven regions conserved among the four Corynebacterium species, as they are amplified under the same conditions. The nucleotide sequences obtained from these seven loci are concatenated, excluding the primer sequences, and then further aligned and compared using the Clustal X program to infer a phylogenetic tree.59 (ii) PCR-Based Analysis  Here, the methods are mainly based on the use of either oligonucleotide sequences from widespread genes and conserved regions of DNA (mostly rRNA genes, but also rpoB genes and others) or specific oligonucleotides that amplify a certain gene or DNA fragment from a particular species that allows its direct identification. An example of the latter is amplification of rRNA intergenic spacer regions or the corynebacterial tox gene. Randomly Amplified Polymorphic DNA (RAPD). PCR is carried out using short primers of random sequences under low-stringency annealing conditions, resulting in the amplification of randomly sized DNA fragments. This method requires carefully controlled conditions in order to obtain good reproducibility. Denaturing Gradient Gel Electrophoresis (DGGE). Specific primers are used to amplify target sequences, frequently from rRNA genes. The PCR-amplified fragments are then separated in a sequence-specific manner by DGGE, yielding a profile of fragments that identify the microbial species under study. In most cases, PCR products of the same length from different species are well separated by DGGE. Amplified Ribosomal DNA Restriction Analysis (ARDRA). This is a sequential method based on PCR amplification of the rRNA gene (frequently 16S rRNA) followed by restriction analysis of the amplified fragment with different restriction enzymes. ARDRA is a reliable and highly

reproducible tool to discriminate very closely related species or isolates of the same species. Multiplex PCR. With this method, more than one locus is simultaneously amplified in the same reaction. The method is suitable for in vivo studies and can be used for strain identification without the need for prior DNA extraction. Target sequences are frequently based on 16S-23S rRNA genes or their intergenic spacer region.

6.1.3  Pathogenesis and Diagnosis 6.1.3.1  Corynebacterium Diphtheriae Corynebacterium diphtheriae is the causative microorganism of diphtheria, although, less often, C. ulcerans is involved. C. diphtheriae species comprise four biotypes (var.): gravis, mitis, intermedius, and belfanti; all of which, with the exception of belfanti, produce the lethal diphtheria exotoxin.60 Diphtheria is usually classified according to its site of manifestation, which most often is the respiratory tract or the skin. C. diphtheriae releases the toxin after colonizing the upper respiratory tract and the skin. Once the toxin is absorbed by the circulatory system, it is distributed to distant organs (myocardium, peripheral nervous system, etc.), where it causes the typical symptoms of the disease. In respiratory diphtheria, the bacteria form a membrane in the posterior structures of the oral cavity (pharynx/larynx). The continued growth of the microorganism on the surface of this membrane generates the characteristic “pseudomembrane,” which reduces airflow from the trachea and thus causes suffocation and possibly death. The genome sequence of C. diphtheriae biotype gravis ribotype “Sankt Petersburg” was published in 2003.61 The bacterial toxin is encoded by a mobile temperate bacteriophage (corynephage) and inhibits protein synthesis in the host cell by ADP-ribosylation of elongation factor 2. However, C. diphtheriae also has a remarkable capacity to cause invasive bloodstream infections even in the absence of toxin production. Systemic complications of C. diphtheriae bacteremia are not unusual and include endocarditis, joint infections, and peripheral embolic disease. The evolutionary origin of nontoxigenic C. diphtheriae is unknown; however, it is very likely that the success of the diphtheria vaccine in preventing toxin-mediated disease resulted in the selection of new pathogenic C. diphtheriae variants expressing other virulence factors (e.g., siderophores or other ironuptake proteins), or equipped with other disease-causing molecular mechanisms that allow the bacteria to overcome host conditions. The increased number of cases of endocarditis caused by these invasive clones testifies to their clinical importance.62 While used in the past, typing systems such as serotyping, phage typing, and bacteriocin typing have largely been replaced by molecular methods—most importantly, Southern hybridization with tox and IS probes, as well as cell wall protein analysis by SDS-PAGE, and gel separation of digested chromosomal DNA.63 For C. diphtheriae, traditional molecular typing methods, including ribotyping, are straightforward and effective in distinguishing isolates of this species.64

66

An interesting molecular approach in the identification of C. diphtheriae strains and subspecies is the above-described multilocus enzyme electrophoresis (MEE). MEE was used to analyze the genetic diversity within a combined panel of preepidemic and epidemic C. diphtheriae strains isolated in 1985–199453 and for further studies; using this technique a clonal group of closely related strains was distinguished, whereas PFGE, RAPD, amplified restriction fragment polymorphism (AFLP), and ribotyping failed to do so. PCR-based methods are faster and simpler, although they sometimes have low discriminatory power and reproducibility. AFLP has been widely used for C. diphtheriae typing and was suggested as a replacement for ribotyping as the “gold standard” of typing methods, because it requires only a small amount of DNA, analyzes the entire genome, and previous sequence information on the target DNA is not necessary. In a recent study, four molecular typing methods (ribotyping, PFGE, RAPD, and AFLP) were used in the characterization of 755 isolates of C. diphtheriae in order to examine the global spread of the pathogen.65 In that study, ribotyping was considered the gold standard for typing due to its discriminatory and reproducible power, followed by RAPD, PFGE, and AFLP. However, RAPD and AFLP are rapid methods that can be used as screening techniques prior to ribotyping, thus omitting the need for complex ribotyping analyses of strains that are unrelated. Polymorphisms in the tox gene and its regulatory element, dtxR, in C. diphtheriae strains isolated in Russia and the Ukraine before and during an epidemic were studied by SSCP analysis,58 while PCR–RFLP analyses of the 16S-23S rRNA internal transcribed spacer differentiated C. diphtheriae from C. ulcerans.66 Publication of the C. diphtheriae genomic sequence enabled a more thorough, precise, and comprehensive search of candidate polymorphic loci for the development of new typing methods of this pathogen. One such method is based on the clustered regularly interspaced short palindromic repeat (CRISPR) technique. CRISPR consists of short sequences (21–37 nucleotides long) that are repeated almost exactly and referred to as “units,” which are separated by similarly sized nonrepetitive sequences called “spacers.”67 CRISPR sequences are present in many of the known bacteria and were advantageously used in hybridization macroarray-based methods to study the internal polymorphism of 156 strains of the epidemic-causing Sankt Petersburg clone in 1997–2002.68 6.1.3.2  C  orynebacterium Species of the Lipophilic Group The majority of the “lipid-requiring,” or “lipophilic” strains of Corynebacterium are readily cultured in the laboratory as they are able to grow on most routinely used culture media, which contain at least minimal amounts of lipids and support the basal growth of lipophilic bacteria. Thus, Corynebacterium grows reasonably well on media enriched with a significant amount of lipids (e.g., 0.1%–1% Tween 80, or 2 drops of sterile serum). Lipophilic corynebacteria are usually fastidious, and slower growing than nonlipophilic

Molecular Detection of Human Bacterial Pathogens

strains. Most are nonhemolytic on blood Columbia agar, contain short corynemycolic acids (C22 to C36), arabinogalactans, and meso-diaminopimelic acid in their cell walls, and produce dehydrogenated menaquinones. In general, they lack the genes required for synthesizing the fatty acids present in the cell membrane and are resistant to a broad range of antibiotics. Corynebacterium urealyticum. This strictly aerobic species was previously included in Corynebacterium group D2 of the Centers for Disease Control (CDC, Atlanta, GA) classification due to its characteristics of very high urealytic activity, nonfermentative, nitrate-negative metabolism, and lipophilicity. However, DNA hybridization analysis invalidated this taxonomic assignment and resulted in the bacterium’s formal designation as C. urealyticum. The description of this new species was also supported by phenotypic tests, such as API profiles, and by chemotaxonomic (presence of fatty acids, isoprenoids, quinones, and phospholipids) and molecular (rRNA gene profiling) analyses.45 The natural habitat of C. urealyticum is the human skin, but it also can be isolated from blood and wounds and readily colonizes the urinary tract. In fact, highly antibiotic-resistant strains of C. urealyticum have been associated with cases of encrusted cystitis and other urinary tract infections in hospital patients.69 C. urealyticum resembles Corynebacterium jeikeium in its colony and cellular morphologies. In addition, the two species have a common habitat and both are remarkably antibiotic resistant. However, both can be clearly distinguished to each other by amplification of the 16S-23S rRNA gene spacer regions.70 Biochemical analyses of C. urealyticum indicate a closest relation to Corynebacterium pseudodiphtheriticum (due to its urealytic activity), although the latter is positive for nitrate reduction and nonlipophilic. The genome sequence of C. urealyticum was recently obtained by pyrosequencing71 and contains about 2020 coding sequences. Of these, 80% are orthologous to the corresponding genes of the Corynebacterium jeikeium genome,72 indicating a close phylogenetic relationship between these two lipid-requiring species. A DNA region with 70 CRISPRs and eight CRISPR-associated genes was identified in the C. urealyticum genome. The DNA repeats are 28 bp in length, separated by variable 33-bp spacer sequences, and similar to the clustered repeats of CRISPRs in the C. jeikeium genome. The presence of C. urealyticum in patients with cystitis has been detected by PCR amplification of a 212-bp band characteristic for this specie.73 Corynebacterium jeikeium. The bacterium is considered part of the normal flora of the human skin, where it predominantly colonizes the axillary, inguinal, and perineal areas. Initially, bacteria from “group JK corynebacteria” (CDC, Georgia At) were associated with cases of bacterial endocarditis that occurred following cardiac surgery and isolated as nondiphtherial corynebacteria from blood cultures. These bacteria were later named C. jeikeium and frequently confirmed as the causative agent of nosocomial infections, especially in immunocompromised patients.

Corynebacterium

Many of the reported C. jeikeium isolates are substantially multiresistant against clinically relevant antibiotics, limiting the therapeutic options in infected patients. Whole-genome sequencing of C. jeikeium showed that its multidrug resistance is encoded by chromosome-associated genes most likely acquired by bacterial horizontal transfer.72 More than 50% of the C. jeikeium genome comprises genes orthologous to those of nonpathogenic corynebacteria, and a closest relation to the C. urealyticum genome has been reported.71,72 The molecular techniques used to identify C. jeikeium are based on species-specific probes for its 16S rRNA.74 Plasmid content and RFLP techniques rRNA-associated have also been used in strain identification, although polymorphisms due to minor changes in RFLP profiles were noted as well.75 Corynebacterium ureicelerivorans. This lipophilic coryneform bacterium generates organic acids from glucose and splits urea very rapidly. C. ureicelerivorans, most likely produces struvite stones, made up of ammonium, magnesium, and phosphates, as has been demonstrated in C. urealyticum and other urea-splitting organisms. C. ureicelerivorans has a close genetic relationship with C. mucifaciens and C. jeikeium. New isolates of C. ureicelerivorans isolated from blood cultures obtained from patients with bacteremia were identified based on phenotypic (API and Biolog systems) and molecular (16S rDNA and rpoB sequencing) characteristics.76 Colonies of these bacteria growing on blood Columbia agar are very small, whitish, dry, and circular, with intact edges. A misidentification of C. ureicelerivorans with C. bovis is possible when only the API system is used, as the identification code is the same for the two species. An equivalent uncertainty likewise results when the Biolog system is used for identification. Instead, molecular typing of C. ureicelerivorans is recommended and is based on rpoB sequence analyses.76 According to sequencing database, the percent similarity of C. ureicelerivorans with C. mucifaciens is very high (91%–95%). Corynebacterium macginleyi. This diphtheroid (named in honor of KJ McGinley, who made important contributions to the study of lipophilic diphtheroids) is oxidase-negative, facultatively anaerobic, and able to produce organic acids from sugars.27 It has been implicated in conjunctivitis and endophthalmitis, but also in endocarditis and septicemia. One of the most interesting reports concerning C. macginleyi is the etiologic agent of keratitis following keratoplasty, described by Suzuki et al.77 In their study, two C. macginleyi isolates were identified by 16S rRNA sequencing (99.2% similarity to the type strain) and further rpoB gene analyses. In the latter, the similarity of the isolates was 97.3% with C. macginleyi and 92% with the closely related C. accolens. Recently, new molecular techniques have been applied in the molecular identification of eye-associated pathogens. For example, multilocus sequence typing (MLST), with seven housekeeping genes (adk, dnaA, fumC, gltA, gyrB, icd, and purA) from sequenced species was used to design consensus primers for further DNA amplification from pathogenic strains. The MLST results were subsequently validated by

67

RAPD analyses, thus confirming the technique’s very good discriminatory power for C. macginleyi and C. accolens.78 Corynebacterium accolens. This species, formerly known as diphtheroid genomospecies II (CDC), is an inhabitant of the upper respiratory tract and is closely related to C. macginleyi. C. accolens was originally characterized by its satellite growth around a Staphylococcus aureus streak on blood agar.8 When grown on Columbia blood agar with lipids, the bacterium forms small, gray, nonhemolytic, and transparent colonies. Many of the reported Corynebacterium-related cases of endocarditis have been due to lipophilic organisms such as C. jeikeium, C. afermentans (subsp. lipophilum), and C. accolens. Like C. macginleyi, which C. accolens most closely resembles, it is negative for maltose fermentation and urease production but positive for glucose fermentation and nitrate reductase production. However, C. accolens can be differentiated from C. macginleyi since the former is negative for alkaline phosphatase. In addition, the two species can be distinguished by ARDRA analyses based on CfoI digestion of amplified 16S rRNA gene fragments.79 6.1.3.3  C  orynebacterium Species of the Nonlipophilic Group Many of the members of this group were easily identified by ARDRA analyses based on their 16S rRNA genes.80 C. xerosis, C. striatum, and C. amycolatum Subgroup. C. xerosis was one of the earliest Corynebacterium species identified as a human pathogen. It exhibits fermentative metabolism and α-glucosidase, alkaline phosphatase, and pyrazinamidase activities. Although this species was previously described as a definite human pathogen, most of the strains tentatively identified in the routine clinical laboratory as C. xerosis have since been reclassified as C. striatum or C. amycolatum. Nowadays, true C. xerosis clinical isolates are extremely rare,6 although we have evidences that isolates obtained from clinical human samples are indeed C. xerosis (LM Mateos, unpublished results). All three species commonly occur in the normal human flora of the skin and the oropharynx, making it difficult to distinguish infection from contamination or colonization. Phenotypically, C. xerosis strains produce yellowish colonies, express mycolic acids, have leucine arylamidase activity, produce lactic acid as a byproduct of glucose fermentation, and are susceptible to the vibriocidal agent O/129. C. amycolatum strains, in contrast, produce whitish-grayish colonies, lack mycolic acids, have either weak or no leucine arylamidase activity, produce large amounts of propionic acid from glucose fermentation, and are mostly resistant to O/129; a culture of C. amycolatum grown on Columbia agar with 5% blood can be seen in Figure 6.1. Accordingly, differentiation between C. amycolatum and C. xerosis is achieved by chemotaxonomic procedures, such as mycolic acid analysis and the detection of propionic acid production from glucose (C. amycolatum), whereas routine identification is difficult because the colonies formed by the two species are very similar in appearance. Macroscopically, C. xerosis

68

Molecular Detection of Human Bacterial Pathogens

of the 16S-23S rRNA intergenic spacer using the enzyme CfoI,82 16S rRNA gene sequencing, and DNA hybridization (% similarity)—must be applied. Polyphasic analyses employing the rpoB and 16S rRNA genes (among others) have been successfully used as well.57

6.2  METHODS 6.2.1  Sample Preparation

FIGURE 6.1  Culture of C. amycolatum grown on Columbia agar with 5% blood.

and C.  striatum colonies can be distinguished because the colonies of C. xerosis are yellowish, dry, and granular, while those of C. striatum are whitish, moist, and smooth. As mentioned earlier, the two species can be differentiated by ARDRA80 and other PCR-derived analyses.55 C. minutissimum. Colonies of this species are macroscopically convex and circular, about 1–1.5 mm in diameter (after 24 h in Columbia agar), with a shiny and moist surface and intact edges. C. minutissimum is the causative agent of erythrasma, an asymptomatic skin infection characterized by scaly, reddish, macular patches in intertriginous areas.81 When tested with the API Coryne strip, C. minutissimum strains may be misidentified as C. jeikeium because the same code number is ascribed to both species; however, the latter requires lipids and is aerobic. C. minutissimum can also be misidentified as C. amycolatum, but certain phenotypic reactions (i.e., DNAase activity), the presence of tuberculostearic acid, and the products obtained from glucose fermentation can help to distinguish between the two. In addition, C. minutissimum is able to grow on medium containing 6.5% NaCl. Molecular analyses have been broadly used, targeting the PCR-amplified 16S rRNA gene. The polymorphism of the divIVA gene (after PCR amplification) has been successfully used to distinguish different nonlipophilic fermentative coryneform species.55 Corynebacterium freneyi. This strain, which is closely related to C. amycolatum and C. xerosis, was first described by Renaud et al.19 Phenotypically, C. freneyi can be differentiated from C. xerosis by its distinct wrinkled colonies, whereas nearly all other routinely applied phenotypic tests do not allow unanimous definitive separation of the two species. Instead, molecular techniques—specifically, RFLP analysis

6.2.1.1  Culture Media for Corynebacterium There are no selective or enrichment procedures that are specifically suitable for Corynebacterium; rather, a wide variety of media can be used for their culture as well as for macroscopic and microscopic studies. Most members of Corynebacterium are able to grow on modified peptoneyeast-glucose media such as Corynebacterium medium (DSMZ Medium 790), containing (per liter): 10.00 g peptone; 1.00 g yeast extract; 2.00 g MgSO4 ∙ 7H2O; and 2.00 g (NH4)2SO4 at pH 7.0. Good complex media include brain heart infusion (BHI, Oxoid), tryptone-soy agar (TSA; Difco), and Columbia agar (Difco) supplemented with 5% horse or sheep blood, with incubation at 35°C–37°C in an air plus 5% CO2-enriched atmosphere (when required). The composition of Columbia agar medium with 5% blood is described below. Chemically defined media are available only for nonpathogenic corynebacteria, with the addition of certain growth factors to stimulate growth (e.g., dihydroxyphenolic compounds such as catechol, and vitamins biotin and thiamine)83; see below for the composition of BMCG medium. Columbia Agar + 5% Sheep Blood (1 L). 5.0 g peptic digest of animal tissue; 5.0 g pancreatic digest of casein; 10.0 g yeast-enriched peptone; 3.0 g pancreatic digest of heart muscle; 1.0 g cornstarch; 5.0 g sodium chloride; 14.0 g agar; 50.0 mL sheep blood; and final pH of 7.3 ± 0.2 at 25°C. Basal Medium for Corynebacterium Growth (Synthe­ tic BMCG) (1 L). 7.0 g (NH4)2SO4; 6.0 g Na2HPO4; 3.0 g KH2P04; 0.5 g NaCl; 1.0 g NH4Cl; 400 mg MgSO4∙7H2O; 20 mg FeSO4∙7H2O; 2 mg MnSO4∙H20; 0.2 mg Na2B4O7∙10H2O; 80 mg (NH4)6Mo7O24∙4H2O; 20 mg ZnSO4∙7H2O; 0.5 mg CuSO4∙5H2O; 15 mg MnCl2∙4H2O; 0.2 mg FeC13∙6H2O; 30 mg biotin; 2 mg thiamine; 3 mg nicotinic acid; 10 µM catechol; and 10 g of a carbon source (frequently glucose). All solutions are adjusted to neutral pH. 6.2.1.2  DNA Isolation This protocol is based on the method described by Vaneechoutte et al.80 Strains are grown on Columbia agar supplemented with 5% (v/v) sheep blood at 37°C (with or without CO2 enrichment). A loopful of the corresponding colony (about 10–20 µL) is suspended in 300 µL of distilled water, boiled for 10 min, agitated thoroughly, and then centrifuged briefly in a benchtop microcentrifuge. The supernatant of the sample preparation is placed in a new Eppendorf tube and is ready to be used for further DNA analyses.

69

Corynebacterium

6.2.1.3  RNA Isolation This protocol is a combination of the RNA extraction methods designed for Trizol (Tri Reagent, Sigma) and silica columns (“Qiagen RNeasy kit,” Qiagen). As complete DNA digestion is crucial to obtain highly pure RNA, the samples are DNase treated twice, first using Turbo DNase (Ambion) and then again with the DNase provided in the column of the RNeasy kit (Qiagen), following the manufacturers’ instructions. In brief, 50 mL of complex liquid medium (TSA or Columbia) is inoculated to reach an optical density at 600 nm (OD600) of 0.05, and the bacteria are harvested at midexponential phase (OD600 = 0.25–0.32). One ml of the culture is transferred to a centrifuge tube and the cells are pelleted at 4000 × g for 5 min at 4°C. The supernatant is discarded, the pellet is mixed with 1 mL of Tri Reagent (Sigma), and the suspension transferred to a 2-mL screw-capped tube containing 500 µL DEPC-treated 0.1-mm zirconia-silica beads. The beads are beat for 2 × 30 s pulses at a speed setting of 6.5 and the tubes allowed to cool at room temperature (RT) for 5 min. After overnight storage at –80°C, the contents of the tubes are thawed on ice and centrifuged for 1 min at 16,100 × g at RT. The supernatant is transferred to a new tube, followed by the addition of 200 µL of chloroform, vigorous shaking for 15 s, and separation of the layers by allowing the tubes to stand for 15 min at RT. After centrifugation at 12,000 × g at 4°C for 15 min, the clear upper aqueous phase (≈ 700 µL) is transferred to a new tube and 500 µL of isopropanol is added. After adequate mixing, the tubes are allowed to stand at RT for 10 min to allow the RNA to precipitate. The precipitated RNA is collected by centrifugation at 12,000 × g at 4°C for 10 min. The supernatant is discarded, the pellet washed with 1.5 mL of 75% ethanol, and the sample centrifuged at 7500 × g at 4°C for 5 min. The supernatant is again discarded and the pellet dried for 10–15 min, resuspended in 90 µL of RNase-free water, and incubated at 65°C for 5 min. Following the addition of 10 µL of Ambion Turbo 10× buffer and 2 µL of Ambion Turbo DNase, the RNA is incubated at 37°C for 30 min and then thoroughly mixed with 350 µL of Qiagen RLT buffer. After the addition of 250 µL of 100% ethanol and thorough mixing, the sample is applied to the RNeasy column. The column is centrifuged for 15 s at 8000 × g, 350 µL of buffer RW1 is then added to the center of the column, and the column centrifuged again for 15 s at 8000 × g. In a separate tube, 10 µL of Qiagen DNase I is gently mixed with 70 µL of RDD buffer. The DNase mix is added to the center of the column, which is then incubated at RT for 15 min. Next, 350 µL of RW1 is loaded onto the center of the column, followed by centrifugation for 15 s at 8000 × g, the addition of 500 µL of buffer RPE to the center of the column, and centrifugation as before. The previous step is repeated, discarding the flow-through, and the sample centrifuged for a further 2 min at 8000 × g. The column is then transferred to a new Eppendorf tube, 50 µL of RNase-free water is added, and the column incubated at RT for 5 min followed by centrifugation for 1 min at 8000 × g. The previous step is repeated, and the RNA quantified at OD260/280 to check for RNA integrity.

6.2.2  Detection Procedures 6.2.2.1  Biolog (GP2) System The Biolog microbial identification system84 is used to analyze gram-positive bacteria (GP2). The test reactions are carried out in a 96-well microtiter plate containing 95 dehydrated substrates. Organisms are prepared by subculturing them in the appropriate growth medium (supplemented with 5% sheep blood or 1% Tween 80 when required). After incubation, the isolates are subcultured on the surface of equivalent solid media. All plates are incubated for 24 h at 35°C under oxic conditions. A bacterial suspension is obtained by removing the bacterial colonies from the surface of the medium and mixing them with 5 mL of 0.85% saline solution. The bacterial suspension is then standardized to 35%–42% transmittance and a 150-μL aliquot of this suspension dispensed into the wells of a Biolog GP2 panel. After the panels have been incubated for 24 h at 35°C, they are read with the Biolog microplate reader (Biolog Microstation 3.5 software and the corresponding database). The Biolog software compares the results obtained with the test strain to the database and provides identification based on distance calculations. 6.2.2.2  API Coryne System Version 2.0 of the API Coryne system85 includes a gallery of 20 microcupules containing dehydrated substrates. The system provides the following biochemical tests: nitrate reduction, pyrazinamidase, pyrrolidonyl arylamidase, alkaline phosphatase, β-glucuronidase, β-galactosidase, α-glucosidase, N-acetyl-β-glucosaminidase, esculin, urease, gelatin hydrolysis, a fermentation (negative) control, glucose, ribose, xylose, mannitol, lactose, sucrose, and glycogen. All isolates should be tested using the procedures recommended by the manufacturer, except for isolates exhibiting lipid dependency. A suspension consisting of colony material from each isolate in 0.3 mL of sterile distilled water is swabbed onto the surface of a TSA agar plate supplemented with 5% sheep blood. For organisms exhibiting lipid dependency, colony material for each isolate is inoculated into 1 mL of TSB (tryptone soy broth, Difco) supplemented with 1% Tween 80, incubated for 4 h at 35°C under oxic conditions, and then spread over the entire surface of a TSA agar plate supplemented with 5% sheep blood. The plates are again incubated for 24 h at 35°C under oxic conditions and the resulting colonies are removed from the surface of the plate and suspended in 3.0 mL of sterile distilled water to a turbidity equivalent to a McFarland standard of 6. The first 11 microcupules of the test-strip gallery are inoculated with this suspension. For the nine microcupules of the fermentation test, a 0.5-mL aliquot of the culture is added to 3.0 mL of GP medium (provided in the system), mixed with a vortex, and the mixture used as inoculant. The urease, negative control, and fermentation-test microcupules are overlaid with sterile oil and all test strips are incubated oxically at 35°C for 24 h. Subsequently, the appropriate reagents are added to each well and the color reactions are read with the provided color chart after a 10-min incubation at RT. The previously determined catalase test results are

70

used as the final test. Reactions are recorded on the provided test-result sheets and a seven-digit octal code is generated. This numerical profile is interpreted with the profile index provided by the manufacturer for a final identification. 6.2.2.3  Identifications Based on 16S rRNAs Genes Sequencing of the 16S ribosomal gene (16S rRNA) is used for confirmatory molecular identification of a test organism. One primer pair is used routinely, with the upper and lower primers consisting of a consensus sequence with respect to specific parts of the amplified gene.54 The size of the anticipated band depends on the primers used for PCR geneamplification analyses. The PCR procedure used to amplify the rDNA sequence is frequently performed as follows: (i) A 1-μL aliquot of a frozen DNA stock (see the method describing total DNA extraction from corynebacteria) is added to 49 μL of a PCR mixture containing 22.7 μL of distilled water, 15.2 μL of 3.3× PCR buffer (provided by the commercial distributor), 4 μL of 10 mM dNTP mix, 1.6 μL of a 25 mM magnesium acetate solution, 2.5 μL of a 10 μM solution of the 5′ (upper) primer, 2.5 μL of a 10 μM solution of the 3′ (lower) primer, and 0.5 μL of the appropriate polymerase (frequently Taq or Pfu); (ii) the PCR amplification is performed in a thermal cycler under the following conditions: an initial 94°C for 1.5 min; 35 cycles of 94°C for 20 s; 55°C for 45 s; 72°C for 4 min; and a final 72°C for 7 min; however, the conditions should be optimized in each case to achieve the best results. The PCR-product mixture for each isolate is then purified using a commercial purification kit. The purified products are sequenced using the Sanger technique, with 3 μL of 1 μM primer, and then PCR-amplified using the previously described program. The completed sequences for each isolate are exported in a readable format and compared to the 16S rRNA gene sequence database using the National Center for Biotechnology Information taxonomy browser. An isolate is considered successfully identified to the species level if similarity with a reference sequence is ≥97%.

6.3  FUTURE PERSPECTIVES AND CONCLUSIONS As corynebacteria constitute part of the normal skin flora, in most cases it is difficult to distinguish between infection, colonization, and contamination by these organisms. Despite this uncertainty, it is important to consider Corynebacterium species as causative organisms of abscesses, especially when these bacteria are isolated as a pure contaminant in samples obtained from different parts of the body or from different isolations of the same patient. The presence of gram-positive bacilli associated with polymorphonuclear cells in the isolated clinical specimen is a strong evidence for a causal role of corynebacteria-mediated disease.6 Pure cultures support a possible disease association, as does the predominance of coryneform bacteria in material from normally sterile sites of the body. Over the last two decades, a much broader range of taxonomic studies of bacteria has gradually replaced classical

Molecular Detection of Human Bacterial Pathogens

techniques based on morphological, physiological, and biochemical characterization. Nowadays, polyphasic taxonomy takes into account all available phenotypic and genotypic data, integrating them in a consensus-oriented classification within the framework of a general phylogeny derived from 16S rRNA gene sequence analysis. In some instances, consensus classification is obtained through the analysis of several parameters (usually phylogenetic ones) that are complemented by phenotypic or additional analyses. Molecular methods provide reliable and highly discriminatory solutions to bacterial characterizations, in contrast to traditional or classical typing techniques. Moreover, new approaches to characterization are being developed. For example, the analysis of large bacterial biomolecules such as proteins, nucleic acids, and sugars by MALDI/MALDI-TOF (mass spectrometry) allows direct identification of the suspected bacterium. Moreover, the technique can also be applied to crude cellular fractions or cellular suspensions, and thus to chemotaxonomic classification. The use of molecular techniques such as PCR amplification in the direct detection of clinical pathogens has several advantages over phenotypic identification. Since a single small colony provides adequate starting material, final identification can be achieved within 8–12 h of culture; furthermore, identification by these approaches is less dependent on culture conditions. The specific detection of microorganisms from clinical samples often requires the use of speciesspecific primers and/or probes (mainly when the analyzed bacterium belongs to a previously sequenced organism). However, the use of universal primers (i.e., primers complementary to well-conserved regions of the bacterial genome) can be applied for specific fragment amplification in most bacterial species, in which case the identification relies on the relative polymorphism of the amplified fragment. Accurate and timely identification of coryneform bacteria causing human infection is warranted for several reasons. Although most of these bacteria are generally of low virulence, the infections they cause may be severe or fatal. Also, the precise identification of coryneform bacterial strains is important because their antimicrobial susceptibilities are variable and an effective antimicrobial therapy optimizes disease outcome. The resistance of corynebacteria to multiple antimicrobial agents has increased over the past several decades and is even characteristic of species such as C. jeikeium, C. urealyticum, and C. amycolatum. For C. diphtheriae, the application of molecular typing/subtyping methods and continuous monitoring of bacterial spread (epidemic, endemic, or imported) can indicate the most effective preventive measures. Molecular-based analyses may also lead to the identification of novel organisms and to the discovery of new associations between established species and human disease. More recently, in silico mining of complete-genome sequences of different coryneform bacteria has resulted in the identification of repeated sequences such as the CRISPR regions of C. diphtheriae68 and multiple minisatellite DNA loci. These and techniques still under development (including

Corynebacterium

those based on newly identified spacers and other such variants) will likely provide the next generation of typing procedures for coryneform bacteria.

ACKNOWLEDGMENTS Research presented in this chapter was funded partially by grants from Junta Castilla y León (LE028A10-2) (2004/120) and Ministerio de Ciencia y Tecnología (BIO2005–02723 and BIO2008–00519) of Spain.

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1. Martín, J.F. et al., Cloning systems in amino acid-producing corynebacteria, Biotechnology, 5, 137, 1987. 2. Lehmann, K.B., and Neumann, R.O. In Lehmann, J.F. (Editor), Atlas und Grundriss der bakteriologie und Lehrbuch der specillen bakteriologischen diagnostik, 1st Ed., p. 1, München, 1896. 3. Goodfellow, M., Suprageneric classification of actinomycetes. In Williams, S.T., Sharpe, M.E., and Holt, J.G. (Editors), Bergey’s Manual of Systematic Bacteriology, p.  2333, Williams and Wilkins, Baltimore, 1989. 4. Collins, M.D., and Cummins, C.S., Genus Corynebacterium. In Sneath, P.H.A., Mair, N.S., Sharpe, M.E., and Holt, J.G. (Editors), Bergey’s Manual of Systematic Bacteriology, Vol. 2, p. 1266, Williams and Wilkins, Baltimore, 1986. 5. Puliti, M. et al., Experimental model of infection with non-toxigenic strains of Corynebacterium diphtheriae and development of septic arthritis, J. Med. Microbiol., 55, 229, 2006. 6. Funke, G. et al., Clinical microbiology of coryneform bacteria, Clin. Microbiol. Rev., 10, 125, 1997. 7. Liebl, W., Corynebacterium nonmedical. In Dworking, M., Falkow, S., Rosemberg, E., Scheifer, K., and Stackebrandt, E. (Editors), The prokaryotes, 3rd Ed., p. 796, Springer, Germany, 2006. 8. Neubauer, M. et al., Corynebacterium accolens sp. nov., a gram-positive rod exhibiting satellitism, from clinical material, Syst. Appl. Microbiol., 14, 46, 1991. 9. Riegel, P. et al., Taxonomic study of Corynebacterium Group ANF-1 strains: Proposal of Corynebacterium afermentans sp.  nov. containing the subspecies C. afermentans subsp. afermentans subsp. nov. and C. afermentans subsp. lipophilum subsp. nov., Int. J. Syst. Bacteriol., 43, 287, 1993. 10. Collins, M.D., Burton, R.A., and Jones, D., Corynebacterium amycolatum sp. nov., a new mycolic acid-less Corynebacterium species from human skin, FEMS Microbiol. Lett., 49, 349, 1988. 11. Riegel, P. et al., Corynebacterium argentoratense sp. nov. from the human throat, Int. J. Syst. Bacteriol., 45, 533, 1995. 12. Yassin, A.F., Steiner, U., and Ludwig, W. Corynebacterium appendicis sp. nov., Int. J. Syst. Evol. Microbiol., 52, 1165, 2002. 13. Hall, V. et al., Corynebacterium atypicum sp. nov. from a human clinical source, does not contain corynomycolic acids, Int. J. Syst. Evol. Microbiol., 53, 1065, 2003. 14. Funke, G., Lawson, P.A., and Collins, M.D., Heterogeneity within human-derived centers for disease control and prevention (CDC) coryneform group ANF-1-like bacteria and description of Corynebacterium auris sp. nov., Int. J. Syst. Bacteriol., 45, 735, 1995.

71 15. Yassin, A.F., Steiner, U., and Ludwig, W., Corynebacterium aurimucosum sp. nov. and emended description of Corynebacterium minutissimum Collins and Jones (1983), Int. J. Syst. Evol. Microbiol., 52, 1001, 2002. 16. Funke, G. et al., Corynebacterium confusum sp. nov., isolated from human clinical specimens, Int. J. Syst. Bacteriol., 48, 1291, 1998. 17. Funke, G., Ramos, C.P., and Collins, M.D., Corynebacterium coyleae sp. nov., isolated from human clinical specimens, Int. J. Syst. Bacteriol., 47, 92, 1997. 18. Riegel, P. et al., Corynebacterium durum sp. nov., from human clinical specimens, Int. J. Syst. Bacteriol., 47, 1107, 1997. 19. Renaud, F.N. et al., Corynebacterium freneyi sp. nov., alphaglucosidase-positive strains related to Corynebacterium xerosis, Int. J. Syst. Evol. Microbiol., 51, 1723, 2001. 20. Furness, G., and Evangelista, A.T., Infection of a nonspecific urethritis patient and his consort with a pathogenic species of nonspecific urethritis Corynebacteria, Corynebacterium genitalium, nov. sp., Invest. Urol., 14, 202, 1976. 21. Funke, G. et al., Corynebacterium glucuronolyticum sp. nov. isolated from male patients with genitourinary infections, Med. Microbiol. Lett., 4, 204, 1995. 22. Renaud, F.N. et al., Corynebacterium hansenii sp. nov., an alphaglucosidase-negative bacterium related to Corynebacterium xerosis, Int. J. Syst. Evol. Microbiol., 57, 1113, 2007. 23. Funke, G. et al., Corynebacterium imitans sp. nov. isolated from patients with suspected diphtheria, J Clin Microbiol., 35, 1978, 1997. 24. Jackman, P.J.H. et al., Classification of corynebacteria associated with endocarditis (group JK) as Corynebacterium jeikeium sp. nov., System. Appl. Microbiol., 9, 83, 1987. 25. Collins, M.D. et al., Corynebacterium kroppenstedtii sp. nov., a novel corynebacterium that does not contain mycolic acids, Int. J. Syst. Bacteriol., 48, 1449, 1988. 26. Funke, G. et al., Corynebacterium lipophiloflavum sp. nov. isolated from a patient with bacterial vaginosis, FEMS Microbiol. Lett., 150, 219, 1997. 27. Riegel, P. et al., Genomic diversity and phylogenetic relationships among lipid-requiring diphtheroids from humans and characterization of Corynebacterium macginleyi sp. nov., Int. J. Syst. Bacteriol., 45, 128, 1995. 28. Merhej, V. et al., Corynebacterium timonense sp. nov. and Corynebacterium massiliense sp. nov., isolated from human blood and human articular hip fluid, Int. J. Syst. Evol. Microbiol., 59, 1953, 2009. 29. Collins, M.D., and Jones, D., Corynebacterium minutissimum sp. nov., nom. rev., Int. J. Syst. Bacteriol., 33, 870, 1983. 30. Funke, G., Lawson, P.A., and Collins, M.D., Corynebacterium mucifaciens sp. nov., an unusual species from human clinical material, Int. J. Syst. Bacteriol., 47, 952, 1997. 31. Aravena-Roman, M. et al., Corynebacterium pilbarense sp. nov., a non-lipophilic species isolated from a Bactec anaerobic vial inoculated with an ankle aspirate, Int. J. Syst. Evol. Microbiol., 60, 1484, 2010. 32. Furness, G., Sambury, S., and Evangelista, A.T., Coryne­ bacterium pseudogenitalium sp. nov. commensals of the human male and female urogenital tracts, Invest. Urol., 16, 292, 1979. 33. Peel, M.M. et al., Human lymphadenitis due to Coryne­ bacterium pseudotuberculosis: Report of ten cases from Australia and review, Clin. Infect. Dis., 24, 185, 1997. 34. Otsuka, Y. et al., Corynebacterium resistens sp. nov., a new multidrug-resistant coryneform bacterium isolated from human infections, J. Clin. Microbiol., 43, 3713, 2005.

72 35. Funke, G., Lawson, P.A., and Collins, M.D., Corynebacterium riegelii sp. nov., an unusual species isolated from female patients with urinary tract infections, J. Clin. Microbiol., 36, 624, 1998. 36. Wattiau, P., Janssens, M., and Wauters, G., Corynebacterium simulans sp. nov., a non-lipophilic, fermentative Coryne­ bacterium, Int. J. Syst. Evol. Microbiol., 50, 347, 2000. 37. Riegel, P. et al., Corynebacterium singulare sp. nov., a new species for urease-positive strains related to Corynebacterium minutissimum, Int. J. Syst. Bacteriol., 47, 1092, 1997. 38. Coyle, M.B., Leonard, R.B., and Nowowiejski, D.J., Pursuit of the Corynebacterium striatum type strain, Int. J. Syst. Bacteriol., 43, 848, 1993. 39. Yassin, A.F., and Siering, C., Corynebacterium sputi sp. nov., isolated from the sputum of a patient with pneumonia, Int. J. Syst. Evol. Microbiol., 58, 2876, 2008. 40. Collins, M.D. et al., Corynebacterium sundsvallense sp. nov., from human clinical specimens, Int. J. Syst. Bacteriol., 49, 361, 1999. 41. Zimmermann, O. et al., Corynebacterium thomssenii sp. nov., a Corynebacterium with N-acetyl-beta-glucosaminidase activity from human clinical specimens, Int. J. Syst. Bacteriol., 48:489, 1998. 42. Feurer, C. et al., Taxonomic characterization of nine strains isolated from clinical and environmental specimens, and proposal of Corynebacterium tuberculostearicum sp. nov., Int. J. Syst. Evol. Microbiol., 54, 1055, 2004. 43. Riegel, P. et al., Isolation of Corynebacterium tuscaniae sp. nov. from blood cultures of a patient with endocarditis, J. Clin. Microbiol., 44, 307, 2006. 44. Riegel, P. et al., Taxonomy of Corynebacterium diphtheriae and related taxa, with recognition of Corynebacterium ulcerans sp. nov. nom. rev., FEMS Microbiol. Lett., 126, 271, 1995. 45. Pitcher, D. et al., Classification of coryneform bacteria associated with human urinary tract infection (group D2) as Corynebacterium urealyticum sp. nov., Int. J. Syst. Bacteriol., 42, 178, 1992. 46. Yassin, A.F., Corynebacterium ureicelerivorans sp. nov., a lipophilic bacterium isolated from blood culture, Int. J. Syst. Evol. Microbiol., 57, 1200, 2007. 47. Funke, G. et al., Multicenter evaluation of the updated and extended API (RAPID) Coryne database 2.0, J. Clin. Microbiol., 35, 3122, 1997. 48. Funke, G., Peters, K., and Aravena-Roman, M., Evaluation of the RapID CB plus system for identification of coryneform bacteria and Listeria spp., J. Clin. Microbiol., 36, 2439, 1998. 49. Adderson, E.E., Boudreaux, J.W., and Hayden R.T., Infections caused by coryneform bacteria in pediatric oncology patients, Pediatr. Infect. Dis. J., 27, 136, 2008. 50. Becker, B. et al., Rapid differentiation between Nocardia and Streptomyces by paper chromatography of whole cell hydrolysates, Appl. Microbiol., 12, 421, 1964. 51. Lechevalier, M. P., Identification of aerobic actinomycetes of clinical importance, J. Lab. Clin. Med., 71, 934, 1968. 52. Minnikin, D.E. et al., Thin-layer chromatography of methanolysates of mycolic acid-containing bacteria, J. Chromatogr., 188, 221, 1980. 53. Popovic, T. et al., Molecular epidemiology of diphtheria in Russia, 1985–1994, J. Infect. Dis., 174, 1064, 1996. 54. Relman, D.A., Universal bacterial 16S rDNA amplification and sequencing. In Persing, D.H., Smith, T.F., Tenover, F.C., and White, T.J. (Editors), Diagnostic Molecular Microbiology;

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Principles and Applications, p. 489, Mayo Foundation/ American Society for Microbiology, Rochester, MN/ Washington, DC, 1993. 55. Letek, M. et al., Identification of the emerging skin pathogen Corynebacterium amycolatum using PCR-amplification of the essential divIVA gene as a target, FEMS Microbiol. Lett., 265, 256, 2006. 56. Khamis, A., Raoult, D., and La Scola, B., rpoB gene sequencing for identification of Corynebacterium species, J. Clin. Microbiol., 42, 3925, 2004. 57. Khamis, A., Raoult, D., and La Scola, B., Comparison between rpoB and 16S rRNA gene sequencing for molecular identification of 168 clinical isolates of Corynebacterium, J.  Clin. Microbiol., 43, 1934, 2005. 58. Nakao, H. et al., Heterogeneity of diphtheria toxin gene, tox, and its regulatory element, dtxR, in Corynebacterium diphtheriae strains causing epidemic diphtheria in Russia and Ukraine, J. Clin. Microbiol., 34, 1711, 1996. 59. Maiden, M.C., Multilocus sequence typing: A portable approach to the identification of clones within populations of pathogenic microorganisms, Proc. Natl. Acad. Sci. USA, 95, 3140, 1998. 60. Efstratiou, A., and George, R.C., Laboratory guidelines for the diagnosis of infections caused by Corynebacterium diphtheriae and C. ulcerans. World Health Organization, Commun. Dis. Public Health, 2, 250, 1999. 61. Cerdeño-Tárraga, A.M. et al., The complete genome sequence and analysis of Corynebacterium diphtheriae CTC13129, Nucleic Acids Res., 31, 6516, 2003. 62. Moreira L.O. et al., Effects of iron limitation on adherence and cell surface carbohydrates of Corynebacterium diphtheriae strains, Appl. Environ. Microbiol., 69, 5907, 2003. 63. Coyle, M.B. et al., The molecular epidemiology of three biotypes of Corynebacterium diphtheriae in the Seattle outbreak, 1972–1982, J. Infect. Dis., 159, 670, 1989. 64. De Zoysa, A. et al., Molecular epidemiology of Coryne­ bacterium diphtheriae from northwestern Russia and surrou­ nding countries studied by using ribotyping and pulsed-field gel electrophoresis, J. Clin. Microbiol., 33, 1080, 1995. 65. De Zoysa, A. et al., Comparison of four molecular typing methods for characterization of Corynebacterium diphtheriae and determination of transcontinental spread of C. diphtheriae based on BstEII rRNA gene profiles, J. Clin. Microbiol., 46, 3626, 2008. 66. Mokrousov, I.V. et al., The genetic typing of Corynebacterium diphtheriae strains by the polymerase chain reaction with universal primers, Zh. Mikrobiol. Epidemiol. Immunobiol., 5, 73, 1996. 67. Jansen, R. et al., Identification of genes that are associated with DNA repeats in prokaryotes, Mol. Microbiol., 43, 1565, 2002. 68. Mokrousov, I., Corynebacterium diphtheriae spoligotyping based on combined use of two CRISPR loci, Biotechnol. J., 2, 901, 2007. 69. Soriano, F. et al., In vitro and in vivo study of stone formation by Corynebacterium group D2 (Corynebacterium urealyticum), J. Clin. Microbiol., 23, 691, 1986. 70. Aubel, D., Renaud, F.N.R., and Freney, J., Genomic diversity of several Corynebacterium species identified by amplification of the 16S-23S rRNA gene spacer regions, Int. J. Syst. Bacteriol., 47, 767, 1997. 71. Tauch, A. et al., The lifestyle of Corynebacterium urealyticum derived from its complete genome sequence established by pyrosequencing, J. Biotechnol., 136, 11, 2008.

Corynebacterium 72. Tauch, A. et al., Complete genome sequence and analysis of the multiresistant nosocomial pathogen Corynebacterium jeikeium K411, a lipid-requiring bacterium of the human skin flora, J. Bacteriol., 187, 4671, 2005. 73. Simoons-Smit, A.M. et al., Chronic cystitis caused by Corynebacterium urealyticum detected by polymerase chain reaction, Eur. J. Clin. Microbiol. Infect. Dis., 19, 949, 2000. 74. Hou, X.G. et al., Genetic identification of members of the genus Corynebacterium at genus and species levels with 16S rDNA-targeted probes, Microbiol. Immunol., 41, 453, 1997. 75. Pitcher, D. et al., An investigation of nosocomial infection with Corynebacterium jeikeium in surgical patients using a ribosomal RNA gene probe, Eur. J. Clin. Microbiol. Infect. Dis., 9, 643, 1990. 76. Fernández-Natal, M.I. et al., Isolation of Corynebacterium ureicelerivorans from normally sterile sites in humans, Eur. J. Clin. Microbiol. Infect. Dis., 28, 677, 2009. 77. Suzuki, T. et al., Suture-related keratitis caused by Coryneb­ acterium macginleyi, J. Clin. Microbiol., 45, 3833, 2007. 78. Eguchi, H. et al., High-level fluoroquinolone resistance in ophthalmic clinical isolates belonging to the species Cory­ nebacterium macginleyi, J. Clin. Microbiol., 46, 527, 2008. 79. Claeys, G. et al., Endocarditis of native aortic and mitral valves due to Corynebacterium accolens: Report of a case

73 and application of phenotypic and genotypic techniques for identification, J. Clin. Microbiol., 34, 1290, 1996. 80. Vaneechoutte, M. et al., Evaluation of the applicability of amplified rDNA-restriction analysis (ARDRA) to identification of species of the genus Corynebacterium, Res. Microbiol., 146, 633, 1995. 81. Darras-Vercambre, S. et al., Photodynamic action of red light for treatment of erythrasma: Preliminary results, Photodermatol. Photo- immunol. Photomed., 22, 153, 2006. 82. Funke, G., and Frodl, R., Comprehensive study of Cory­ nebacterium freneyi strains and extended and emended description of Corynebacterium freneyi Renaud, Aubel, Riegel, Meugnier, and Bollet 2001, J. Clin. Microbiol., 46, 638, 2008. 83. Liebl, W. et al., High efficiency electroporation of intact Corynebacterium glutamicum cells, FEMS Microbiol. Lett., 53, 299, 1989. 84. Lindenmann, K., von Graevenitz, A., and Funke, G., Evaluation of the Biolog system for the identification of asporogenous, aerobic gram-positive rods, Med. Microbiol. Lett., 4, 287, 1995. 85. Funke, G., and Bernard, K.A., Coryneform gram-positive rods. In Baron, P.R., Jorgensen, E.J., Landry, J.H., and Pfaller, M.A. (Editors), Murray Manual of Clinical Microbiology, 9th Ed., p. 485, American Society for Microbiology, Washington, DC, 2007.

7 Cryptobacterium Futoshi Nakazawa CONTENTS 7.1 Introduction........................................................................................................................................................................ 75 7.1.1 Classification and Morphology............................................................................................................................... 75 7.1.2 Biology, Habitat, and Lifestyle............................................................................................................................... 75 7.1.3 Genetic and Genomic Features............................................................................................................................... 76 7.1.4 Clinical Features and Pathogenesis........................................................................................................................ 77 7.1.5 Diagnosis................................................................................................................................................................ 77 7.2 Methods.............................................................................................................................................................................. 78 7.2.1 Sample Preparation................................................................................................................................................. 78 7.2.2 Detection Procedures.............................................................................................................................................. 78 7.3 Conclusion and Future Perspectives................................................................................................................................... 79 References.................................................................................................................................................................................... 79

7.1  INTRODUCTION 7.1.1  Classification and Morphology The genus Cryptobacterium was proposed by Nakazawa et al. in 1999 for strains of asaccharolytic, anaerobic, short gram-positive rods.1 Initially, these strains were isolated from human oral cavities as Eubacterium-like organisms. The genus Eubacterium has been defined mainly on the basis of negative metabolic characteristics.2 Because of its broad definition, a large number of novel gram-positive rods isolated from oral cavities have been classified previously as members of the genus Eubacterium. Therefore, it is inevitable that considerable heterogeneity exists among the species within the genus. In fact, it is described that the genus Eubacterium contains many gram-positive rods that are phylogenetically unrelated, and some of which qualify as members of the genus Eubacterium but could not be assigned to any of the named species.2–8 In addition, several novel genera have been proposed for some of asaccharolytic, anaerobic, short grampositive rods (e.g., Cryptobacterium, Slackia, Eggerthella, and Mogibacterium), and some species of Eubacterium have been transferred to the novel genera according to their phylogenetic characteristics.1,9,10 Cryptobacterium curturn is the sole species in the genus Cryptobacterium on the basis of morphological and biochemical characteristics, SDS-PAGE of whole-cell protein and Western immunoblotting analysis, G + C content, DNA–DNA hybridization, and 16S rRNA gene sequence data as member of the family Coriobacteriaceae.1 To date, the family Coriobacteriaceae is defined only in terms of the 16S rRNA gene signature nucleotide, just like the subclass Coriobacteridae and the order Coriobacteriales.11 Indeed, it would seem appropriate to

examine the value of chemotaxonomy in improving the definition of the order and family. At the time of writing, the family Coriobacteriaceae proposed by Stackebrandt et al. comprises the following 13 genera: Adlercreutzia, Asaccharobacter, Atopobium, Collinsella, Coriobacterium, Cryptobacterium, Denitrobacterium, Eggerthella, Enterorhabdus, Gordonibacter, Olsenella, Paraeggerthella, and Slackia.1,10,12–23 Cryptobacterium cells are very short (0.4 × 0.8–1.0 μm), obligate anaerobic, nonmotile, nonspore forming rods that occur singly or in masses. The growing cells are grampositive, but occasionally cells in older cultures are gramnegative. Transmission electron micrographs of thin sections of cell of C. curtum 12–3T (=ATCC 700863) showed a singlelayer cell wall approximately 10 nm in thickness (Figure 7.1), which is typical cell wall of gram-positive bacteria. Pili- or flagella-like structures were not found. The scanning electron micrograph of C. curtum 12–3T, occurring in chains, is also available.24 On BHI-blood agar plates, Cryptobacterium cells form minute, circular, convex, and translucent colonies of less than 1 mm in diameter (about 0.3–0.5 mm) without hemolysis, even after prolonged incubation in an anaerobic glove box. Growth in broth media is very poor with or without carbohydrates. Also, it is described that C. curtum strain DSM 15641 (=strain 12–3T) can also be grown anaerobically in DSMZ medium 78 (Chopped Meat Medium).24,25

7.1.2  Biology, Habitat, and Lifestyle Strains of the genus Cryptobacterium are strictly anaerobic and asaccharolytic rods. They do not ferment adonitol, amygdalin, arabinose, cellobiose, erythritol, esculin, fructose, 75

76

Molecular Detection of Human Bacterial Pathogens

asaccharolytic strains. There are no chemotaxonomic data available to C. curtum strain 12–3T.

100 nm

CW CM

FIGURE 7.1  Transmission electron microphotograph of a thin section of cell of C. curtum 12–3T (=ATCC 700863T) showing the cytoplasmic membrane (CM) and typical gram-positive cell wall (CW).

galactose, glucose, glycogen, inositol, lactose, maltose, mannitol, mannose, melezitose, melibiose, rhamnose, ribose, salicin, sorbitol, starch, sucrose, trehalose, or xylose, and are inert in most conventional biochemical tests. They are positive for arginine hydrolysis. Nitrate reduction is negative and they do not hydrolyze esculin, starch, or gelatin. In addition, they do not produce indole, urease, or catalase. They also do not produce any metabolic end product in peptone-yeast extract-glucose broth. C. curtum strains 12–3T and KV43-B were isolated from periodontal pocket of a human adult with periodontal disease and necrotic dental pulp, respectively.8,26 It has been suggested that C. curtum is one of the causative species identified by nonculture techniques in the microbial flora of an acute dental abscess with other anaerobic gram-positive rods such as Eubacterium sulci, Mogibacterium timidum, and Mogibacterium vescum.27 C. curtum and the closely related species such as Slackia exigua do not utilize carbohydrates, and are very difficult to cultivate. Optimal doubling time is 12 h.28 Uematsu et al. described that amino acids are important metabolic substrates for growth of these asaccharolytic strains, especially arginine and lysine.29,30 And they also reported that C. curtum metabolize arginine as source of energy via the arginine deiminase pathway and produce substantial amounts of citrulline and ornithine. Arginine may be provided by the enzymatic degradation of peptides, especially in the inflamed lesions where trypsin-like enzymes produce peptides containing arginine at the C-terminal. In this sense, it would appear that these infectious lesions, such as periodontal pocket, necrotic dental pulp, and dental abscess, have suitable environments for the Cryptobacterium together with other anaerobic and

7.1.3  Genetic and Genomic Features As reported previously, the G + C contents of oral isolates that are asaccharolytic, anaerobic, and short gram-positive rods range from 38 to 62 mol%, as determined by HPLC.1,3,5,6 Similarly, the G + C content of C. curtum 12–3T and KV43-B are 50 and 51 mol%, respectively, which are at the same level as Mogibacterium pumilum (type species of the genus Mogibacterium) or Eubacterium limosum (type species of the genus Eubacterium). The DNA–DNA relatedness data are very useful for identification of asaccharolytic, anaerobic, and gram-positive rods from human clinical specimens.31 In fact, the DNA– DNA hybridization studies with the membrane filter method indicated that the DNA of C. curtum strain 12–3T exhibited a high level (97% homology) of reassociation with the DNA of strain KV43-B. Moreover, the DNA from strains 12–3T and KV43-B exhibited very low levels (1%–11% homology) of relatedness to type species of the other genera that are asaccharolytic, strictly anaerobic, and short gram-positive rods.1,31 The 16S rRNA gene sequence of C. curtum strain 12–3T (1429 bases in length) was registered as accession number AB019260 in GenBank.1 According to the evolutionary tree constructed with the 16S rRNA gene sequence of C. curtum strain 12–3T and other sequences of the related gram-positive bacteria selected from the GenBank database by the sequence similarity search using the BLAST and FASTA algorithms, the strain 12–3T and KV43-B can be distinguished from the asaccharolytic Eubacterium species, including the type species of Eubacterium, E. limosum. These phylogenetic data completely agreed with previous results obtained by RFLP analysis of 16S rRNA.8 Furthermore, although strain 12–3T had a close phylogenetic relationship with Coriobacterium glomerans and Denitrobacterium detoxzjicans, the sequence data showed distinct lines exhibiting specific phylogenetic association with C. glomerans or D. detoxijicans.1 These phylogenetic studies also indicated that strain 12–3T and KV43-B belonged to a large group of the Actinobacteria, and form a cluster with the genus Atopobium, Coriobacterium, and Denitrobacterium (Figure 7.2). Other studies of the 16S rRNA gene sequences data also demonstrated that strain 12–3T formed one cluster with the genus Eggerthella, Collinsella, Slackia, Adlercreutzia, Asaccharobacter, Denitrobacterium, Slackia, and Coriobacterium, all of which are members of the family Coriobacteriaceae.24,31 The strain 12–3T (= DSM 15641 = ATCC 700683 = CCUG 43107) is the type strain of C. curtum, which is the sole species within the genus Cryptobacterium. It is interesting that C. curtum 12–3T was initially isolated from human periodontal pocket as an asaccharolytic Eubacterium-like strain that is a member in the phylum Firmicutes. On the basis of a polyphasic taxonomic study—that notably uses 16S rRNA

77

Cryptobacterium

0.02 Knuc 100

Atopobium fossor JCM 9981T Atopobium minutum ATCC 33267T Eggerthella lenta ATCC 25559T Cryptobacterium curtum ATCC 700683T

100 98

Collinsella aerofaciens JCM 10188T Coriobacterium glomerans DSM 20642T

90

Slackia exigua ATCC 700122T 99

infections such as head and neck infections, abdominal infection bacteremia associated with malignancy, and genital tract infection associated with intrauterine devices.7,46–55 The published reports suggest that Cryptobacterium species may play some roles in oral pathological processes as important pathogens, like other bacterial species of asaccharolytic, anaerobic, and gram-positive rods in human hosts. In animal hosts, no data are currently available on pathogenesis of infections caused by Cryptobacterium species. Specific virulence factors remain unknown. The availability of the complete genome sequences of C. curtum strain 12–3T should give new insights about the virulence of this microorganism.24

Slackia heliotrinireducens ATCC 29202T

FIGURE 7.2  Evolutionary tree based on 16S rRNA gene sequence comparison showing the phylogenetic position of C. curtum 12–3T (=ATCC 700863T). The dendrogram was created by using the neighbor-joining method. The numbers on the tree indicate bootstrap values (%) calculated from 1000 replications.

gene sequence and DNA–DNA hybridization data—C. curtum 12–3T showed very distant and isolated position within the family Coriobacteriaceae in the phylogenetic tree constructed with the 16S rRNA gene sequences. It was finally replaced as a species in the genus Cryptobacterium, family Coriobacteriaceae, phylum Actinobacteria.1,24 The whole genome sequence of C. curtum 12–3T was reported in 2009 as the first complete genome sequence of the family Coriobacteriaceae.24 According to the report, the genome size is 1,617,804 bp in length and comprises one main circular chromosome with a 50.9% DNA G + C content. Of the 1422 predicted genes, 1364 are protein-coding genes, 58 are RNA genes, and 1117 of these genes encode proteins with known function.24

7.1.4  Clinical Features and Pathogenesis C. curtum was described as opportunistic pathogen with a typical occurrence in the oral cavity, and that is involved in such oral infections as periodontitis, inflammations, and abscesses.24 C. curtum strain 12–3T and VK43-B were initially isolated as the closest related rods to the genus Eubacterium taxonomically because of its strict anaerobic, asaccharolytic, and grampositive rod.1 In the same way, the numerous bacterial strains of asaccharolytic, anaerobic, and gram-positive rods isolated from human oral cavities have been classified as members of Eubacterium because of the broad definition of the genus. Furthermore, many of these strains have been documented as aetiologic agents of chronic periodontitis on the basis of their frequent isolation from diseased periodontal sites.32–41 In addition, sera of patients with periodontitis show higher antibody titers against some of these species than those of healthy people, suggesting that these bacteria cause immunological reactions in periodontal lesions.42–45 Moreover, they have been recognized as pathogens in oral and dental infections such as advanced caries, pulpal infections, and dentoalveolar abscesses, and also in nonoral, clinically significant

7.1.5  Diagnosis Conventional Techniques. Strictly anaerobic conditions are required for the growth of the Cryptobacterium species. In our laboratory, Cryptobacterium strains are cultured in an anaerobic glove box (model AZ-Hard, Hirasawa, Japan) containing 80% N2, 10% H2, and 10% CO2. Moreover, to ensure strictly anaerobic conditions in the box, the reduction of methyl viologen (at –446 mV in oxidation reduction potential) is carefully checked when culture procedures are carried out. Even under these anaerobic conditions, the growth of Cryptobacterium strains in broth media is very poor with or without carbohydrates because they are asaccharolytic and difficult to culture. Although no enrichment method and no specific selective medium have been reported for growth of Cryptobacterium strains, arginine and citrulline support the growth of the microorganisms. Cryptobacterium strains utilize arginine via the arginine deiminase pathway and produce substantial amounts of citrulline and ornithine.28 The type strain 12–3T of C. curtum was isolated on brain heart infusion-yeast extract blood (sheep) agar (BHI-blood agar) plates at 37°C for 3 days in an anaerobic glove box.8 Many other agar media or broths can be used for cultivation and isolation of Cryptobacterium, as Kavrommatis et al. demonstrated the growth of C. curtum on DSMZ medium 78.24,25 Also, Wade et al. maintained the Slackia species, which are closely related to Cryptobacterium, on Fastidious Anaerobe Agar supplemented with 5% horse blood under anaerobic conditions.10 In the routine analysis of human oral samples, Cryptobacterium isolates are recognized on BHI-blood agar at 37°C for 5–7 days. C. curtum strain 12–3T and KV43-B yield nonpigmented, nonhemolytic tiny colonies on this medium. After 96 h of incubation at 37°C in the anaerobic glove box, the colonies are 0.3–0.5 mm in diameter, and circular, convex, and whitish with a smooth surface.1 Members of the genus Cryptobacterium are nonmotile and nonsporulating. Other biochemical tests are mainly negative for these microorganisms, except arginine hydrolysis, which is positive. Cryptobacterium strains show neither urease nor catalase activity. Nitrate reduction, gelatin and starch liquefaction, and esculin hydrolysis are negative. Indole is generally not produced. Although no an end product from peptone-yeast extract-glucose broth, C. curtum produces

78

ornithine from BHI broth media with the addition of arginine or citrulline. Using rapid ID 32 A system, Cryptobacterium strains display the following positive enzymatic reactions: arginine arylamidase, leucyl glycine arylamidase, leucine arylamidase, alanine arylamidase, glycine arylamidase, and histidine arylamidase activities. Several of the previously cited characteristics are variable among strains of asaccharolytic species making phenotypic identification difficult. Phenotypic identification of anaerobic and asaccharolytic gram-positive rods from human oral samples is mainly based on negative fermentation characteristics, which allow the identification of the genus Eubacterium mainly encountered in human clinical samples. In fact, strain 12–3T and KV43-B were initially wrongly placed in the genus Eubacterium, and then relocated in the genus Cryptobacterium of the family Coriobacteriaceae. Strictly anaerobic grampositive rods, which are asaccharolytic, have no end products, and are negative in most of the conventional biochemical tests, are often submitted to molecular identification. Molecular Techniques. Molecular methods for identification of Cryptobacterium species include DNA–DNA hybridization and 16S rRNA gene sequencing, which are performed on DNA extracts obtained by Marmur protocol with slight modification.3,56 The two strains of C. curtum each showed 97 mol% in homology of whole DNA relatedness. But they showed less than 11 mol% in the homologies against other strains in the family Coriobacteriaceae. Furthermore, these strains formed a cluster together with members of the family Coriobacteriaceae on the basis of their 16S rRNA gene sequences. The 16S rRNA gene sequence comparisons showed that C. curtum has a close phylogenetic relationship with Denitrobacterium detoxzjicans and Coriobacterium glomerans. But these sequence similarities were 88.5% with D. detoxzjicans and 86.6% with C. glomerans. The phylogenetic analysis indicated that C. curtum was clearly distinct from the other genera in the family Coriobacteriaceae. Molecular techniques for taxonomic characterization of Cryptobacterium species particularly genomic structure analysis, are not detailed in this chapter but may be found in the following reference.10,24

7.2  METHODS 7.2.1  Sample Preparation Cellular morphology is determined by examining cells grown on BHI-blood agar plates. Biochemical characteristics are tested by the procedures described previously by Holdeman et al.25 No end products from peptone-yeast extract-glucose broth have been confirmed by gas chromatography.57 Whole-cell protein profiles are examined by sodium dodecyl sulfate-poly acrylamide gel electrophoresis as described previously.58 After these proteins are transferred to a nitrocellulose membrane, an immune rabbit antiserum is used as the first antibody and followed by incubation with goat antirabbit immunoglobulin G conjugated with

Molecular Detection of Human Bacterial Pathogens

peroxidase as the second antibody in the Western immunoblotting analysis.59 A Rapid ID 32A kit (bioMérieux) is used for enzymic profile determination as recommended by the manufacturer. DNA is isolated and purified by following a modification of the Marmur protocol.3,56 Commercialized kits, such as the InstaGene DNA purification Matrix (BIO-RAD Laboratories), are also available for genomic DNA extraction. The G + C content of DNA is determined by high-performance liquid chromatography, as described previously.60 DNA–DNA hybridization is performed by the membrane filter method according to the procedures described by Meyer et al., with multiprime DNA labeling kit (Amersham Pharmacia Biotech) and32 P-dCTP.61

7.2.2  Detection Procedures Cryptobacterium could be identified by sequencing of the 16S rRNA gene and phylogenetic analysis. Also the data of complete genome sequence of C. curtum type strain may be useful for identification of Cryptobacterium. Procedure 1. The 16S rRNA gene is amplified by PCR with a nucleotide primer set [27 forward primer; 5′-AGA GTT TGA TCM TGG CTC AG-3′, located at positions 8–27 (Escherichia coli numbering), and 1492 reverse primer; 3′-TTC AGC ATT GTT CCA TYG GCAT-5′, located at positions 1492–1513 (E. coli numbering)]. The PCR reaction is performed with Premix Taq (Takara-bio, Japan) according to the manufacturer’s instructions in a thermal controller (e.g., model PTC-100; MJ Research, MA) using the following parameters: 35 cycles 94°C for 1 min, 45°C for 2 min, and 72°C for 2 min. 2. Amplification products are separated by 2.0% agarose gel electrophoresis and visualized by staining with ethidium bromide. According to these conditions, the amplification products of 1.5 kb are obtained from the agarose gel for 16S rRNA gene. 3. For sequence of 16S rRNA from the PCR, a Thermo Sequenase Fluorescent Labeled Primer Cycle Sequencing Kit (Amersham Pharmacia Biotech) is used with the 11 universal primers labeled with Cy-5, following the manufacturer’s protocol. The sequences of the 16S rRNA are analyzed with a DNA sequencer (e.g., ALFexpress, Amersham Pharmacia Biotech, Buckinghamshire). 4. The sequences are aligned with each other and connected by using SEQMAN II of the LASERGENE program (DNASTAR). Programs MEGALIGN of LASERGENE, CLUSTAL W, and NJPLOT are used to compare sequences and to construct an evolutionary tree by the neighbor-joining method. Also, confidence intervals are assessed by NJPLOT and CLUSTAL W with bootstrap analysis.

79

Cryptobacterium

Note. Matsuki et al. designed 16S rRNA gene-targeted groupspecific primers for the member of the Atopobium cluster that includes C. curtum. These primers may be useful for rough identification of this microorganisms.62



7.3  CONCLUSION AND FUTURE PERSPECTIVES



Cryptobacterium species is isolated from human oral cavities and may act as opportunistic pathogens in infectious lesions. Only C. curtum, a type species of the genus Cryptobacterium, is known at the present time. The two strains of C. curtum, strain 12–3T isolated from periodontal pocket and strain KV43-B isolated from necrotic dental pulp, can be differentiated from strains of other genera in the family Coriobacteriaceae by phenotypic characteristics including arginine hydrolysis, nitrate reduction, API ZYM code in Rapid ID32A system, protein profile of whole bacterial cell, and Western immunoblotting pattern. 16S rRNA gene sequencing is known as a useful tool for identification of the genus and the species levels, and is frequently used to identify such anaerobic gram-positive rods from human oral cavities. In fact, according to the results of 16S rRNA gene sequence, Cryptobacterium is established as genus in the family Coriobacteriaceae, which contains anaerobic, asaccharolytic, and gram-positive rods. Restriction fragment-length polymorphism analysis of the amplified 16S rRNA using the endonucleases AluI and SmaI is also a convenient method of distinguishing Cryptobacterium from other genera. Furthermore, DNA–DNA hybridization studies allow the genus Cryptobacterium to be distinguished from closely related taxa such as the genera Collinsella, Eggerthella, and Slackia. The genus Cryptobacterium, with C. curtum as the type species, is identified on the basis of a polyphasic taxonomic study, notably by using 16S rRNA gene sequence and DNA–DNA hybridization data. Little is known about the role of Cryptobacterium species in human oral cavities. With the genome sequence of type species of the genus Cryptobacterium having been recently completed, further details regarding virulence factors or metabolism for the bacterium will be uncovered, contributing to increased knowledge of the roles of Cryptobacterium in human oral diseases.

REFERENCES

1. Nakazawa, F. et al., Cryptobacterium curtum gen. nov., sp. nov., a new genus of gram-positive anaerobic rod isolated from human oral cavities, Int. J. Syst. Bacteriol., 49, 1193, 1999. 2. Moore, W.E.C., and Moore, L.V.H., Genus Eubacterium. In Sneath, P.H.A. et al. (Editors), Bergey’s Manual of Systematic Bacteriology, Williams and Wilkins, Baltimore/London, 1986. 3. Nakazawa, F., and Hoshino, E., Genetic relationships among Eubacterium species, Int. J. Syst. Bacteriol., 44, 787, 1994. 4. Cheeseman, S. et al., Phylogeny of oral asaccharolytic Eubacterium species determined by 16S ribosomal DNA sequence comparison and proposal of Eubacterium infrmun sp. nov. and Eubacterium tardum sp. nov, Int. J. Syst. Bacteriol., 46, 957, 1996.





















5. Poco, S.E. et al., Eubacterium exiguum sp. nov., isolated from human oral lesions, Int. J. Syst. Bacteriol., 46, 1120, 1996. 6. Poco, S.E. et al., Eubacterium minutum sp. nov., isolated from human periodontal pockets, Int. J. Syst. Bacteriol., 46, 31, 1996. 7. Wade, W.G. et al., An unclassified Eubacterium taxon in acute dentoalveolar abscess, J. Med. Microbiol., 40, 115, 1994. 8. Sato, T. et al., Restriction fragment-length polymorphism analysis of 16S rDNA from oral asaccharolytic Eubacterium species amplified by polymerase chain reaction, Oral. Microbiol. Immunol., 13, 23, 1993. 9. Nakazawa, F. et al., Description of Mogibacterium pumilum gen. nov., sp. nov. and Mogibacterium vescum gen. nov., sp. nov., and reclassification of Eubacterium timidum (Holdeman et al. 1980) as Mogibacterium timidum gen. nov., comb. nov, Int. J. Syst. Evol. Microbiol., 50, 679, 2000. 10. Wade, W.G. et al., The family Coriobacteriaceae: Reclassification of Eubacterium exiguum (Poco et al. 1996) and Peptostreptococcus heliotrinreducens (Lanigan 1976) as Slackia exigua gen. nov., comb. nov. and Slackia heliotrinreducens gen. nov., comb. nov., and Eubacterium lentum (Prevot 1938) as Eggerthella lenta gen. nov., comb. nov., Int. J. Syst. Bacteriol., 49, 595, 1999. 11. Stackebrandt, E. et al., Phylogenetic analysis of the genus Desulfotomaculum: Evidence for the misclassification of Desulfotomaculum guttoideum and description of Desul­ fotomaculum orientis as Desulfosporosinus orientis gen. nov., comb. nov., Int. J. Syst. Bacteriol., 47, 1134, 1997. 12. Stackebrandt, E. et al., Proposal for a new hierarchic classification system, Actinobacteria classis nov., Int. J. Syst. Bacteriol., 47, 479, 1997. 13. Naruo, T. et al., Adlercreutzia equolifaciens gen. nov., sp. nov., an equol-producing bacterium isolated from human faeces, and emended description of the genus Eggerthella, Int. J. Syst. Evol. Microbiol., 58, 1221, 2008. 14. Minamida, K. et al., Asaccharobacter celatus gen. nov., sp. nov., isolated from rat caecum, Int. J. Syst. Evol. Microbiol., 58, 1238, 2008. 15. Collins, M.D., and Wallbanks, S., Comparative sequence analyses of the 16S rRNA genes of Lactobacillus minutus, Lactobacillus rimae and Streptococcus parvulus: Proposal for the creation of a new genus Atopobium, FEMS Microbiol. Lett., 74, 235, 1992. 16. Kageyama, A., and Benno, Y., Emendation of genus Collinsella and proposal of Collinsella stercoris sp. nov. and Collinsella intestinalis sp. nov., Int. J. Syst. Evol. Microbiol., 50, 1767, 2000. 17. Haas, F., and König, H., Coriobacterium glomerans gen. nov., sp. nov. from the intestinal tract of the red soldier bug, Int. J. Syst. Bacteriol., 38, 382, 1988. 18. Anderson, R.C. et al., Denitrobacterium detoxificans gen. nov., sp. nov., a ruminal bacterium that respires on nitrocompounds, Int. J. Syst. Evol. Microbiol., 50, 633, 2000. 19. Kageyama, A. et al., Phylogenetic evidence for the transfer of Eubacterium lentum to the genus Eggerthella as Eggerthella lenta gen. nov., comb. nov., Int. J. Syst. Bacteriol., 49, 1725, 1999. 20. Clavel, T. et al., Enterorhabdus caecimuris sp. nov., a novel member of the family Coriobacteriaceae isolated from a mouse model of spontaneous colitis, Int. J. Syst. Evol. Microbiol., in Press, 2010. 21. Clavel, T. et al., Isolation of bacteria from the ileal mucosa of TNFdelta ARE mice and description of Enterorhabdus mucosicola gen. nov., sp. nov., Int. J. Syst. Evol. Microbiol., 59, 1805, 2009.

80 22. Wurdemann, D. et al., Gordonibacter pamelaeae gen. nov., sp. nov., a new member of the Coriobacteriaceae isolated from a patient with Crohn’s disease, and reclassification of Eggerthella hongkongensis Lau et al. 2006 as Paraeggerthella hongkongensis gen. nov., comb. nov., Int. J. Syst. Evol. Microbiol., 59, 1405, 2009. 23. Dewhirst, F.E. et al., Characterization of novel human oral isolates and cloned 16S rDNA sequences that fall in the family Coriobacteriaceae: description of Olsenella gen. nov., reclassification of Lactobacillus uli as Olsenella uli comb. nov. and description of Olsenella profusa sp. nov., Int. J. Syst. Evol. Microbiol., 51, 1797, 2001. 24. Mavrommatis, C. et al., Complete genome sequence  of Cryptobacterium curtum type strain (12–3T), Standards Genom. Sci., 1, 93, 2009. 25. Holdeman, L.V., Cato, E.P., and Moore, W.E.C., Anaerobe Laboratory Manual, 4th Ed., Blacksburg, VA, Virginia Polytechnic Institute and State University, 1977. 26. Sundqvist, G., Bacteriological studies of necrotic dental pulps, Umei University Dissertation, 7, 1, 1976. 27. Robertson, D., and Smith, A.J., The microbiology of the acute dental abscess, J. Med. Microbiol., 58, 155, 2009. 28. Uematsu, H. et al., Degradation of arginine by Slackia exigua ATCC 700122 and Cryptobacterium curtum ATCC 700683, Oral Microbiol. Immunol., 21, 381, 2006. 29. Uematsu, H., and Hoshino, E., Degradation of arginine and other amino acids by Eubacterium nodatum ATCC 33099, Microbiol. Ecol. Health Dis., 9, 305, 1996. 30. Uematsu, H. et al., Degradation of arginine and other amino acids by butyrate-producing asaccharolytic anaerobic gram-positive rods in periodontal pockets, Arch. Oral. Biol., 48, 423, 2003. 31. Nakazawa, F., and Hoshino, E., DNA–DNA relatedness and phylogenetic positions of Slackia exigua, Slackia heliotrinireducens, Eggerthella lenta, and other related bacteria, Oral Microbiol. Immunol., 19, 1, 2004. 32. Hill, G.B. et al., Characteristics and sites of infection of Eubacterium nodatum, Eubacterium timidum, Eubacterium brachy, and other asaccharolytic eubacteria, J. Clin. Microbiol., 25, 1540, 1987. 33. Holdeman, L.V. et al., Description of Eubacterium timidum sp. nov., Eubacterium brachy sp. nov. and Eubacterium nodatum sp. nov., isolated from human periodontitis, Int. J. Syst. Bacteriol., 30, 163, 1980. 34. Martin, S.A. et al., A comparison of the reactivity of Eubacterium species with localized and serum immunoglobulin from rapidly progressive and adult periodontitis, J. Periodontol., 59, 32, 1986. 35. Moore, W.E.C. et al., Bacteriology of severe periodontitis in young adults, Infect. Immun., 38, 1137, 1982. 36. Moore, W.E.C. et al., Bacteriology of moderate (chronic) periodontitis in mature adult humans, Infect. Immun., 42, 510, 1983. 37. Moore, W.E.C. et al., Comparative bacteriology of juvenile periodontitis, Infect. Immun., 48, 507, 1985. 38. Slayne, M.A., and Wade, W.G., The humoral response to asaccharolytic Eubacterium species in periodontitis, Microbiol. Ecol. Health. Dis., 7, 283, 1994. 39. Uematsu, H., and Hoshino, E., Predominant obligate anaerobes in human periodontal pockets, J. Periodontal. Res., 27, 15, 1992. 40. Uematsu, H. et al., Eubacterium saphenum sp. nov., isolated from the human periodontal pockets, Int. J. Syst. Bacteriol., 43, 302, 1993.

Molecular Detection of Human Bacterial Pathogens 41. Wade, W.G. et al., The effects of antimicrobial acrylic strips on the subgingival microfora in chronic periodontitis, J. Clin. Periodontol., 19, 127, 1992. 42. Gunsolley, J.C. et al., Serum antibodies to periodontal bacteria, J. Periodontol., 61, 412, 1990. 43. Tew, J.G. et al., Serum antibody reactive with predominant organisms in the subgingival flora of young adults with generalized severe periodontitis, Infect. Immun., 48, 303, 1985. 44. Tolo, K., and Jorkjend, L., Serum antibodies and loss of periodontal bone in patients with rheumatoid arthritis, J. Clin. Periodontol., 17, 288, 1990. 45. Tolo, K., and Schenck, K., Activity of serum immunoglobulin G, A and M to six anaerobic oral bacteria in diagnosis of periodontitis, J. Periodontal. Res., 20, 113, 1985. 46. Ando, N., and Hoshino, E., Predominant obligate anaerobes invading the deep layers of root canal dentine, Int. Endod. J., 23, 20, 1990. 47. Edwardsson, S., Bacteriological studies on deep areas of carious dentine, Odontol. Revy. Suppl., 32, 1, 1974. 48. Hoshino, E. et al., Bacterial invasion of non-exposed pulp, Int. Endod. J., 25, 2, 1992. 49. Hoshino, E., Predominant obligate anaerobes in human carious dentin, J. Dent. Res., 64, 1195, 1985. 50. Sato, T. et al., Predominant obligate anaerobes in necrotic pulps of human deciduous teeth, Microbiol. Ecol. Health Dis., 6, 269, 1993. 51. Brook, I., and Frazier, E.H., Significant recovery of nonsporulating anaerobic rods from clinical specimens, Clin. Infect. Dis., 16, 476, 1993. 52. Fanstein, V. et al., Bacteremia caused by non-sporulating anaerobes in cancer patients, Medicine (Baltimore), 68, 151, 1989. 53. Brook, I., Anaerobic bacterial bacteremia: 12-year experience in two military hospitals, J. Infect. Dis., 160, 1071, 1983. 54. Backer, E.D. et al., Antibiotic susceptibility of Atopobium vaginae. BMC Infect. Dis., 6, 51, 2006. 55. Goldstein, E.J.C. et al., In vitro activities of daptomycin, vancomycin, quinupristin-dalfopristin, linezolid, and five other antimicrobials against 307 gram-positive anaerobic and 31 Corynebacterium clinical isolates. Antimicrob. Agents Chemother., 47, 337, 2003. 56. Marmur, J., A procedure for the isolation of DNA from microorganisms, J. Mol Biol., 3, 208, 1961. 57. Hoshino, E., and Sato, M., Production and degradation of formate by Veillonella dispar ATCC 17745, J. Dent. Res., 65, 903, 1986. 58. Laemmli, U.K., Cleavage of structural proteins during the assembly of the head of bacteriophage T4, Nature, 227, 68, 1970. 59. Nakazawa, F., and Hoshino, E., Immunological specificity of oral Eubacterium species, J. Gen. Microbiol., 39, 2635, 1993. 60. Katayama-Fujimura, Y. et al., Estimation of DNA base composition by high performance liquid chromatography of its nuclease PI hydrolysate, Agric. Biol. Chem., 48, 3169, 1984. 61. Meyer, S.A., and Schleifer, K.H., Deoxyribonucleic acid reassociation in the classification of coagulase positive Staphylococci, Arch. Microbiol., 7, 183, 1978. 62. Matsuki, T. et al., Use of 16S rRNA gene-targeted groupspecific primers for real-time PCR analysis of predominant bacteria in human feces, Appl. Environ. Microbiol., 70, 7220, 2004.

8 Gardnerella Rita Verhelst, Hans Verstraelen, Piet Cools, Guido Lopes dos Santos Santiago, Marleen Temmerman, and Mario Vaneechoutte CONTENTS 8.1 Introduction........................................................................................................................................................................ 81 8.1.1 Cell Wall Structure and Chemical Composition.................................................................................................... 81 8.1.2 Taxonomy............................................................................................................................................................... 82 8.1.3 Biotypes.................................................................................................................................................................. 82 8.1.4 Pathogenesis: Virulence Factors............................................................................................................................. 82 8.1.5 Clinical Features..................................................................................................................................................... 83 8.1.6 Epidemiology.......................................................................................................................................................... 84 8.2 Methods.............................................................................................................................................................................. 85 8.2.1 Phenotypic Identification........................................................................................................................................ 85 8.2.2 Genotypic Identification......................................................................................................................................... 85 8.2.2.1 Sample Preparation.................................................................................................................................. 86 8.2.2.2 Detection Procedures............................................................................................................................... 86 8.3 Conclusions and Future Perspectives................................................................................................................................. 89 References.................................................................................................................................................................................... 90

8.1  INTRODUCTION The genus Gardnerella, consisting of a single facultatively anaerobic Gram-variable rod species, Gardnerella vaginalis, is classified under the family Bifidobacteriaceae, suborder Actinomycineae, phylum Actinobacteria. Although the bacterium possesses a thin gram-positive cell wall, it may appear either gram-positive or gram-negative under the microscope. The clinical significance of G. vaginalis, as well as its taxonomic position, has long been a matter of dispute largely due to variable Gram staining and lack of appropriate detection and identification methods (reviewed in Catlin1). Thus far, more than 1000 papers were addressed to this small (0.4 by 1.0–1.5 µm), pleomorphic, rod-shaped, fastidious bacterium organism. Almost a century ago, Curtis et al.2 reported that Gramstained purulent vaginal discharges contained mainly gramnegative bacilli that he and his colleagues were unable to cultivate. It was not until 1953 that Leopold3 succeeded in isolating such bacteria from men with prostatitis and women with cervicitis. The organism was named Haemophilus vaginalis by Gardner and Dukes in 19554 because it was a gram-negative, rod-shaped bacterium that could be isolated on blood agar and it was believed to be responsible for a characteristic vaginal discharge.

8.1.1  Cell Wall Structure and Chemical Composition Based on electron microscopy data, Reyn et al.5 concluded that, although the organism does not retain crystal violet

during conventional Gram-staining procedures, the thin walls and septa of osmium-fixed G. vaginalis resemble those of gram-positive bacteria. Harper and Davis6 and O’Donnell et al.7 confirmed the gram-positive nature of the cell wall, based on the presence of the amino acids alanine, glutamic acid, glycine and lysine, and on the absence of the dibasic amino acid diaminopimelic acid (DAP) in the peptidoglycan. Also, based on electron microscopy studies, Sadhu et al.8 concluded that the cell wall unequivocally has a grampositive type of organization and that the unusual thinness of the cell walls (8–12 nm) accounts for the gram-negative staining tendency of G. vaginalis. Further evidence that G.  vaginalis most properly groups with the gram-positive bacteria came from the study of Gelber et al.,9 who found that this species expresses vaginolysin, a hemolysin belonging to the cholesterol-dependent cytolysins that have only been described in gram-positive organisms so far. However, evidence for a gram-negative cell wall composition was gathered as well. Criswell et al.10,11 argued for a gram-negative cell wall on the basis of the presence of a wide range of amino acids and because the cell wall has a trilaminar structure, consisting of two electron-dense layers separated by an electron-translucent layer and the presence of only low amounts of peptidoglycan (i.e., only 20% of the total cell wall weight). However, these studies have been faulted because the amino acid profiles for their control organisms were at variance with the currently accepted profiles.6,7 Also, Greenwood and Pickett12 interpreted the lamellar cell wall 81

82

structure as more closely resembling that of gram-negative organisms and identified a lipopolysaccharide (LPS)-like cell wall component. However, the absence of DAP and hydroxy fatty acids, as well as the almost complete absence of endotoxin by the Limulus amebocyte lysate assay, indicate that G. vaginalis does not contain classical LPS.6–8,13,14 In addition, studies on antibiotic susceptibility led to contradicting conclusions on the nature of the cell wall. Although the antibiotic susceptibility pattern of G. vaginalis resembles that of gram-positive bacteria according to Reyn et al.5 and Piot et al.,15 Kharsany et al.16 claimed that G. vaginalis is neither typically gram-positive nor typically gram-negative since antimicrobial agents regarded as specifically active against gram-positive or gram-negative organisms showed relatively poor activity. It should be mentioned that another important gram-positive member of disturbed vaginal microflora (i.e., Lactobacillus iners) typically stains gram-negative and that, also because the similarities of their colony morphology, L. iners has been misidentified on several occasions as G. vaginalis (unpublished data). At present, on the basis of taxonomic data (see below), it is clear that G. vaginalis should be considered a gram-positive bacterium.

8.1.2  Taxonomy Haemophilus vaginalis was renamed to Corynebacterium vaginale by Zinneman and Turner,17 based on the absence of requirements for X and/or V factors (hemin and NAD, respectively), which are needed for the growth of accepted Haemophilus species, the tendency to retain the crystal violet dye in the Gram reaction, and a corynebacterium-like morphology. Reyn et al.5 compared H. vaginalis with four possibly related species that had shown a similar degree of Gram-variability and suggested that H. vaginalis should be included in Corynebacterium or Butyribacterium, rather than in Lactobacillus and that this organism may be closely related to the genus Propionibacterium. However, based on volatile acid production and fermentation end product analysis, Moss and Dunkelberg18 showed that this species did belong to neither the genera Corynebacterium, Butyribacterium nor Lactobacillus. Three large taxonomic studies published in 198012,15 and 199619 analyzed data obtained by a variety of biochemical methods, DNA–DNA hybridization, 16S rRNA sequence analysis, and electron microscopy. Greenwood and Pickett12 found that “Corynebacterium vaginale” formed a new taxon and proposed the name Gardnerella vaginalis, defining the organism as a facultative anaerobic, rod-shaped, catalaseand oxidase-negative, gram-negative to Gram-variable bacterium with laminated cell walls that produces acetic acid as the major end product of fermentation of carbohydrates. In the same year, Piot et al.15 supported the creation of a new genus, but found little ground for G. vaginalis to be related to established gram-negative taxa. Pointing at differences in Gram-stain appearance depending on the growth condition and on the age of the culture, they found that G. vaginalis had

Molecular Detection of Human Bacterial Pathogens

an antibiotic susceptible pattern as found in gram-positive bacteria. Further phylogenetic analysis by Van Esbroeck et al.,19 based on 16S rRNA gene sequence analysis, placed G. vaginalis, still the sole member of the genus Gardnerella, in the gram-positive family Bifidobacteriales. G. vaginalis is indeed closely related to the genus Bifidobacterium, with the highest level of similarity to the type species Bifidobacterium bifidum (i.e., 93.1%) and exhibits fructose-6-phosphate phosphoketolase activity, a feature that was assumed to be specific for the genus Bifidobacterium.20 However, the difference in GC-content between G. vaginalis (42 mol%) and bifidobacteria (55–67 mol%),21 is too large to consider G. vaginalis as a genuine Bifidobacterium species.

8.1.3  Biotypes On the basis of the measurement of the activities of three enzymes (i.e., β-galactosidase, lipase, and hippurate hydrolase), G. vaginalis isolates were assigned to eight different biotypes by Piot et al.22 Another scheme obtained by adding fermentation of arabinose, galactose, and xylose, revealed respectively 17 and 33 different biotypes in two studies.23,24 Based on the latter scheme, different G. vaginalis biotypes were found in women with and without bacterial vaginosis.25 Although, based on the scheme of Piot et al., the association of specific biotypes with bacterial vaginosis remains controversial until today. When the distributions of different G. vaginalis biotypes from women with and without bacterial vaginosis in three countries were examined, biotypes 1, 2, and 5 were found to be the most predominant groups present at all locations.22 However, in a longitudinal study, G. vaginalis strains of biotypes 1, 5, and 6 were found to be most common in all clinical situations.25 Of the 261 isolates examined in that study, the lipase-positive group, consisting of biotypes 1, 2, 3, and 4, was suggested to be more prevalent among women with bacterial vaginosis, a finding confirmed by Numanovic´ et al.26 Aroutcheva et al.27 found diversity in the bacterial vaginosis group and a predominance of biotype 5 in women with normal vaginal microflora. Recently, Moncla et al.28 pointed at methodological differences in the previous studies and argued that 4-methylumbelliferyl-oleate and other 4-methylumbelliferyl-derivatives should not be used for the detection of bacterial lipase activity, and urged to reconsider possible associations between bacterial vaginosis and certain biotypes.

8.1.4  Pathogenesis: Virulence Factors Several factors have been shown to contribute to the ability of bacteria to colonize and infect the host epithelium. Pili. A key feature to vaginal colonization is the ability of G. vaginalis to adhere to the vaginal epithelium. Patterson et al.29 recently compared bacterial adherence to vaginal epithelial cells, biofilm-producing capacities, and cytotoxic activity of bacterial vaginosis-associated organisms, suggesting that G. vaginalis is more virulent than many of the bacterial species frequently isolated from bacterial vaginosis.

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G. vaginalis is undoubtedly equipped with multiple virulence properties and consequently, the idea that it is merely an initial colonizer that paves the way for additional species is being revisited.9,29–31 Bacterial adherence is clinically typically reflected by the presence of clue cells on microscopic examination of a vaginal smear. It was recently convincingly confirmed by FISH that clue cells are vaginal squamous epithelial cells coated with G. vaginalis and other anaerobic bacteria causing bacterial vaginosis.31 Clue cells were first described by Gardner and Dukes in 1955 and were named as such because these cells provide an important “clue” to the diagnosis of bacterial vaginosis.4 At the ultrastructural level, G. vaginalis adherence is thought to relate to the presence of pili extending from the cell wall and to the presence of an exopolysaccharide (see below). Johnson and Davies32 documented by electron microscopy that at least some G. vaginalis strains carry narrow pili with a thickness between 3.0 and 7.5 nm. Boustouller et al. subsequently showed by electron microscopy that the pili were present among G. vaginalis strains freshly isolated from clinical specimens, whereas the expression of these pili tended to disappear upon repeated culture.33 Accordingly, Fredricsson et al.34 found that in vitro, G. vaginalis showed insufficient adherence to vaginal epithelial cells to produce so-called clue cells. Exopolysaccharide. G. vaginalis has been shown to produce an electron-dense, web-like, fibrillar exopolys­ accharide,35 which was found to confer adherence of G. vaginalis cells to vaginal epithelial cells.36 Vaginolysin. G. vaginalis produces a cytolysin, a 60 kDa protein that is a member of the cholesterol-dependent cytolysin family of toxins, and that recognizes the complement regulatory molecule CD59 on the surface of human cells.9,37 This species-specific lysis of human cells has limited mechanistic studies of bacterial vaginosis and its adverse consequences by the absence of a suitable animal model. Vaginolysin is a pore-forming exotoxin that lyses endothelial cells, neutrophils, and erythrocytes (and therefore previously was called a hemolysin).38–41 The lysis of neutrophils might explain the relative absence of neutrophils in bacterial vaginosis. Furthermore, vaginolysin triggers the immune system, induces interleukin-8 production by human epithelial cells, and a specific secretory immunoglobulin A targeting the G. vaginalis vaginolysin has been shown to be upregulated in the setting of bacterial vaginosis.42,43 Sialidase and Prolidase Activity. G. vaginalis produces sialidase and prolidase, two hydrolytic enzymes that may have a role in degrading several key mucosal protective factors, such as mucins, as well as contributing to exfoliation and detachment of vaginal epithelial cells.44 Phospholipase A2 Activity. G. vaginalis has phospholipase A2 activity, and hence may be directly implicated in the initiation of preterm labor through the conversion of arachidonic acid to prostaglandins xxx.45

Iron Acquisition. As iron is an essential growth factor, but also plays an important role in the virulence potential of many bacterial pathogens, a number of bacteria have developed high-affinity iron acquisition mechanisms. Basically, three types of mechanisms have been described (i.e., the secretion of high-affinity iron chelators or siderophores, the expression of cell-surface receptors that directly bind to iron-containing compounds, and hemolysis).46 In G. vaginalis, there is evidence that all three mechanisms are present, including the production of siderophores,46 the binding of iron carriers like catalase,47 lactoferrin,48 transferrin,48 heme,49 and hemoglobin,50 and the production of the aforementioned vaginolysin leading to hemolysis with release of iron-rich compounds from red blood cells. Biofilm Formation. New insights in the pathogenic role of G. vaginalis came from fluorescent in situ hybridization (FISH) studies on vaginal biopsies. Biopsies collected from women with and without bacterial vaginosis were hybridized with a variety of bacterial rRNA-targeted probes.31 Typically, subjects with bacterial vaginosis had an adherent biofilm that was primarily composed with three bacterial groups, of which G. vaginalis was most prominent. G. vaginalis constituted 60%–90% of the biofilm mass, Atopobium accounted for 1%–40% of the biofilm mass, and lactobacilli were present between 1% and 5% in only 20% of the biopsy samples. While this adherent bacterial biofilm was present in 90% of bacterial vaginosis patients, only 10% of women with a normal vaginal microflora had a similar biofilm. Patterson et al.51 have highlighted the physiological differences of G. vaginalis bacteria grown under different conditions. In their study, planktonic G. vaginalis were more sensitive to hydrogen peroxide (5-fold) and lactic acid (4–8-fold) than G. vaginalis biofilm bacteria. Furthermore, the recognition that G. vaginalis forms vaginal biofilm provides a plausible explanation for the persistency and recurrence of bacterial vaginosis in many women, since bacteria in biofilms are more resistant to antibiotics compared to planktonic organisms.52–55 Saunders et al.56 investigated the potential of lactobacilli to attempt clearance of the G. vaginalis biofilm. G. vaginalis biofilms grown in vitro could be displaced by the probiotic Lactobacillus reuteri RC-14 and to a limited extent by Lactobacillus iners, a nonhydrogen peroxide-producing Lactobacillus commonly found in the vaginal econiche, especially in disturbed vaginal microflora.57,58 Future studies on the structure and composition of biofilms will further improve our understanding of the pathogenic role of G. vaginalis in bacterial vaginosis.

8.1.5  Clinical Features The most common infection caused by G. vaginalis is bacterial vaginosis in women. Bacterial vaginosis is a disruption of vaginal microflora characterized by a decrease in Lactobacillus species and increase in the quantity of anaerobic bacteria. G. vaginalis is present in up to 95% of cases of bacterial vaginosis1 and nearly all women with bacterial

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vaginosis have high concentrations of G. vaginalis.59–62 About half of all women microscopically confirmed bacterial vaginosis will present symptoms—the remainder being completely asymptomatic. When symptoms are present, women will present most typically with profuse, off-white, thin, and homogeneous discharge, quite different from the thick, white, curdy discharge observed with vaginal candidiasis. Vaginal discharge with bacterial vaginosis is typically accompanied by an unpleasant, fishy odor. This characteristic vaginal discharge is actually the only symptom—if present at all—and other vulvovaginal complaints such as soreness, pain, or itch, as often present with vulvovaginal candidiasis, are usually not associated with bacterial vaginosis. Similarly, on inspection of the vulvar area and the vaginal walls, no additional signs will be found, as bacterial vaginosis is not associated with (appreciable) vaginal inflammation, and hence no erythema or swelling occur. In men, genital infection associated with G. vaginalis presents as balanoposthitis, which may be accompanied by diffuse erythema and pruritus of the glans, coronal sulcus, and meatus urethrae.63,64 Urinary tract infection with G. vaginalis in men has also been described as urethritis or urethral syndrome.65–68 During pregnancy, G. vaginalis can be involved in upper genital tract infection, including chorioamnionitis and amniotic fluid infection,69,70 infections associated with adverse pregnancy outcome such as late fetal loss, stillbirth, and preterm birth. Procedures during delivery may also be complicated by G. vaginalis infection, such as wound infection following episiotomy71 and following caesarean section,72 and bacteremia occurring during caesarean section.73 Vertical transmission to the neonate can occur, and neonatal infections with G. vaginalis have been anecdotic reported, including G. vaginalis-infected scalp hematoma associated with electronic fetal monitoring,74,75 neonatal conjunctivitis,76 and disseminated neonatal infection presenting as meningitis77 and neonatal sepsis.78,79 Sepsis and abscesses in multiple organ sites with G. vaginalis have also been described in immunocompetent men and women.80–82 Several case reports have also pointed at joint and musculoskeletal infections with G. vaginalis, including vertebral osteomyelitis and discitis,83,84 and hip arthritis.85 G. vaginalis has also been associated with several autoimmune disorders, including reactive arthritis86,87 and Reiter’s syndrome.88

8.1.6  Epidemiology The epidemiology of G. vaginalis in practice partly translates to the epidemiology of bacterial vaginosis. Bacterial vaginosis involves the overgrowth of G. vaginalis as a virtually absolute finding. Among children and prepubertal girls bacterial vaginosis is rather rare, except in case of sexual abuse, even though vaginal carriage of G. vaginalis identified through PCR is

Molecular Detection of Human Bacterial Pathogens

common in young girls.89 Swidsinski et al.90 investigated G. vaginalis carriage in 50 premenarchal girls by FISH and found that dispersed growth of Gardnerella occurred in 10%, whereas none of the 50 girls showed cohesive Gardnerella. Higher rates of G. vaginalis have been found in sexually abused girls (7%–19%) as compared to nonabused girls (1%– 4.2%) in most91–93—though not all—studies.94 Once beyond the menarche however, bacterial vaginosis is observed even among adolescent girls and sexually inexperienced girls, and young women in general, at an appreciable frequency, though at significantly lower rates as compared to sexually active adolescents and young women.95,96 Shafer et al. reported that up to one-third of nonsexually active adolescents harbored G. vaginalis, compared to 60% of sexually active adolescents. Another culture-based study of 120 asymptomatic 14- to 17-year-old females found a twofold higher prevalence of G. vaginalis (34%) in sexually active females compared to women who reported no penetrative sexual activity (17%), albeit a nonsignificant difference. The largest sample up to date involved U.S. women entering the military, with a mean age of 19.1 years, and found bacterial vaginosis in women reporting to never have had vaginal intercourse to occur at a rate of 18%, whereas their sexually experienced counterparts had bacterial vaginosis at a rate of 28%.97 In adult, sexually experienced women, G. vaginalis is a common constituent of the normal vaginal microflora, albeit present in low numbers when compared to the predominant Lactobacillus species. In a European population, G. vaginalis was detected among pregnant women with normal vaginal microflora through species-specific PCR at a rate of 28.9%.98 In a U.S. study involving women attending an STD clinic, G. vaginalis was detected with normal vaginal microflora through species-specific PCR at a rate of 58.7%.99 Women with lesbian orientation, or generally womenwho-have-sex-with-women (WSW), present rather consistently with very high rates of bacterial vaginosis.100–102 Weyers et al.103 characterized the microflora of the penile skin-lined neovagina of transsexual women and found that, although it is difficult to establish to what extent a bacterial vaginosis-like condition is present among these transsexual women, G. vaginalis was present in 30% of the women. Carriage of G. vaginalis is also common in men. The two largest cohort studies on male carriage of G. vaginalis conducted until present—both involving male attendees of a sexually transmitted disease clinic—documented male urethral carriage of G. vaginalis at a rate of 11.4% (49/430) in the United Kingdom104 and of 4.5% (10/309) in Sweden.105 Male carriage of G. vaginalis may even be higher than estimated from the aforementioned studies as urethral sampling may not be the optimal approach to document it. Kinghorn et al.63 found a significantly higher rate of G. vaginalis isolation from preputial than from urethral swabs. Swidsinski et al.106 recently made a similar observation, reporting that desquamated epithelial cells loaded with G. vaginalis could only reliably be recovered in a urine specimen if the praeputium was not pulled back during voiding. These authors

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documented among 100 men admitted to a department of internal medicine that dispersed Gardnerella was found in 4% of males, while cohesive Gardnerella was present in 7%. Remarkably, using PCR, Schwebke et al.107 found no difference in the prevalence rates of G. vaginalis between male sexual partners of women with and without bacterial vaginosis. However, this unexpected finding may be the result of the fact that both groups of men were recruited from the same high-risk cohort of patients attending an STD clinic. In addition, several culture-based studies have also documented the presence of G. vaginalis in semen samples108–113 —possibly pointing at a seminal or prostatic reservoir—whereby in one study G. vaginalis was recovered from semen in as much as 38% of 58 men attending an infertility clinic.109 Evidence to sustain the theory that this microorganism can affect sperm characteristics, thereby causing infertility, was not found.114 However, although G. vaginalis does not affect semen quality and does not cause an inflammatory reaction in infertile men, seminal colonization by this microorganism should be considered clinically relevant, since it may be transmitted to the female genital tract through coitus.109,112,113,115 Overall, G. vaginalis is closely linked to sexual behavior and activity, though it remains unclear through which mechanisms sexual activity enhances G. vaginalis carriage and colonisation.89,116,117 Mechanistic studies of bacterial vaginosis and its adverse consequences have been limited by the absence of suitable animal models since G. vaginalis has been isolated from the reproductive tract of mares only.118 Interestingly, Piot et al.22 found that biotypes isolated from women with bacterial vaginosis and from the urethras of their sexual partners were identical in 90% of the couples. Briselden et al.25 found that women who acquire bacterial vaginosis are more likely to have a shift in biotype than women who had normal microflora at followup, suggesting that the G. vaginalis isolates recovered from women who develop bacterial vaginosis represent newly acquired strains rather than overgrowth of previously colonizing biotypes. Furthermore, Simoes et al.30 found that G. vaginalis biotype 1 strains might be responsible for a higher rate of sexual transmission of HIV, since 50% of these strains were able to induce HIV expression in U1 cells.

8.2  METHODS 8.2.1  Phenotypic Identification Several media possessing differential and/or selective features can be used to isolate G. vaginalis from clinical specimens (reviewed in Catlin1).119,120 At present, human blood-bilayer-Tween (HBT) agar, developed by Totten et al.,121 is the most satisfactory differential selective medium. It consists of a bottom layer of Columbia colistin-nalidixic acid agar, as described by Golberg and Washington,119 supplemented with Proteose Peptone no. 3, amphotericin B, and Tween 80, and a top layer of the same composition to which 5% human blood is added. Often, minimal criteria

such as the characteristics of colonies surrounded by zones of clear hemolysis with diffuse edges on HBT medium, the typical appearance of Gram-stained cells and a negative catalase test are used for the presumptive identification of G. vaginalis from vaginal fluid.122 However, it should be noticed that HBT agar fails to inhibit a variety of gram-positive bacteria, such as lactobacilli, coryneforms, and streptococci.122,123 We found that 3 out of 10 hemolytic strains isolated on HBT agar were identified through sequencing of the 16S rRNA gene as L. iners (unpublished data). Since L. iners strains are oxidase and catalase-negative, positive for β-galactosidase and hippurate hydrolase, and variable in their lipase activity, these strains may be misidentified as G. vaginalis biotypes 1 or 6. Moreover, as for G. vaginalis, short gram-positive L. iners rods can have a gram-negative stain appearance.61 Misidentification of L. iners as G. vaginalis might have led to erroneous conclusions on the role of these species in bacterial vaginosis. Ison et al.124 found that the addition of gentamicin (4 mg/L) instead of colistin to HBT medium increases the inhibition of non-Gardnerella species. Rapid conventional methods for detection and identification of G. vaginalis were reviewed by Catlin.1 It should be noted that the API-ZYM system for enzyme detection,125 as well as the API-20A and Minitek systems,126 were considered unreliable for the identification of G. vaginalis in the diagnostic laboratory. Also, several papers report the failure to detect G. vaginalis bacteremia using blood culture media containing the anticoagulant sodium polyanethole sulphate (SPS).69,127–130 Reimer et al. have suggested that the incidence of bacteraemia caused by G. vaginalis may be higher than the number of reported cases since SPS, routinely added to most blood culture media, inhibits growth of G. vaginalis unless gelatin is also present.131 To tackle the limitations of conventional methods, molecular methods are increasingly used to identify microbes in clinical samples. However, one should realize that molecular methods also experience technical problems. For example, PCR amplification can fail due to the presence of PCR inhibitors. Ironically, initial attempts at amplification of the bacterial 16S rRNA gene from inoculated blood culture media failed due to the presence of SPS, the component that caused growth inhibition of G. vaginalis in SPS containing culture media.132 Appropriate DNA extraction methods to remove PCR inhibitors are presented in the section “sample preparation.” In addition, care should be taken not to miss certain species when “universal” primers are used, as discussed below (see section on sequencing).

8.2.2  Genotypic Identification Material for identification of G. vaginalis can be obtained by self-swabbing133–136 or after placement of a nonlubrificated speculum.57 The swabs need to be rotated against the vaginal wall at the midportion of the vault and carefully removed to prevent contamination with the vulva and introitus microflora.

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If culture media cannot be directly inoculated, the swabs should be placed in a transport medium. Swabs on solid transport medium are not preferred when performing direct molecular identification methods since the transport medium might cause PCR inhibition (unpublished observations). Flocked swabs are preferred for the purpose of direct molecular identification. The use of an ESwab (Copan, Brescia, It) with liquid Amies transport medium has the advantage that the same swab can be used for the purpose of Gram stain, culture, and PCR.137 Alternatively, a flocked swab can be stored dry in an empty tube when frozen at –20°C within 5 h after collection.61,138 Using dry swabs avoid problems of incompatibility of commercial collection kits and transport media for multiple testing of specimens with different nucleic acid amplification tests.139 8.2.2.1  Sample Preparation Crude DNA Extracts of Cultured Isolates. Crude DNA extracts from cultured strains can be obtained by simple alkaline lysis as follows—resuspend one bacterial colony in 20 µL of lysis buffer (0.25% sodium dodecyl sulfate, 0.05 N NaOH) and heat at 95°C for 15 min. Spin down drops that are present on the lid and dilute the cell lysate by adding 180 µL of distilled water. Freeze at –20°C and use the supernatant for PCR analysis. DNA Extracts from Clinical Samples. G. vaginalis are gram-positive bacteria, for which most conventional extraction methods have a relatively poor recovery of nucleic acids.140 In order to detect or accurately quantify G. vaginalis in biological samples, DNA extraction requires chemical lysis, use of lytic enzymes, or physical disruption of the gram-positive cell wall. Several studies have reported promising results for bead beating to optimize isolation of microbial DNA.140,141 Also, the muramidases lysozyme and mutanolysin, enzymes that degrade the gram-positive peptidoglycan layer, have been shown to improve DNA recovery from gram-positives.57,142 In addition, proteinase K can be used to break down contaminating proteins and nucleases that are liberated during lysis of several bacteria. After this pretreatment step, DNA can be extracted using commercially available kits. However, a common limitation to PCR-based methods is failed amplification due to the presence of inhibitory substances in the sample. PCR inhibitors include heme compounds found in blood,143 aqueous and vitreous humors of the eye,144 heparin,145 EDTA, urine,146 and polyamines.147 To deal with this problem, PCR inhibitors must be diluted, inactivated, or removed from the sample using an appropriate extraction method for each specific sample type. For example, Fredricks and Relman found that an organic extraction procedure with benzyl alcohol and guanidine hydrochloride buffer successfully removes SPS, yielding DNA that can be amplified by PCR without further processing or dilution.132 Here, we provide three examples of sample preparation methods for the detection and identification of G. vaginalis from clinical samples.

Molecular Detection of Human Bacterial Pathogens

(i) Vaginal dry swabs and manual DNA extraction method57,61  For DNA extraction from the dry vaginal swabs, the QIAamp DNA mini kit (Qiagen, Hilden, Germany) is used, according to the manufacturer’s recommendations, with minor modifications. The dry swab specimen is swirled for 15 s in 400 µL of lysis buffer (20 mM Tris-HCl, pH 8.0; 2 mM EDTA; 1.2% Triton). Fifty units of mutanolysin (25 U/ µL) (Sigma, Bornem, Belgium) are added and the samples are incubated for 30 min at 37°C. After the addition of 20 µL Proteinase K (20 mg/mL) and 200 µL AL buffer (Qiagen), samples are incubated for 30 min at 56°C. Next, 200 µL of ethanol is added and DNA is purified by adding the lysate to the Qiagen columns, as described by the manufacturer. Finally, the total bacterial DNA is eluted with 100 µL of AE buffer (Qiagen). DNA extracts are stored at –20°C. (ii) Vaginal ESwab and semiautomated DNA extraction method  Pretreatment of samples is carried out by adding 200 µL of buffer (20 mM Tris-HCl, pH 8.3, 0.5% SDS) and 2 µL  of mutanolysin (25U/µL) to 200 µL of ESwab medium, followed by incubation for 15 min at 37°C. After adding 10 µL of proteinase K solution (25 mg/mL), the mixture is further incubated for 15 min at 55°C. The pretreated samples are stored at –80°C until further processing on the NucliSENS® easyMAGTM semiautomated DNA extraction automate (bioMérieux). Total bacterial DNA is eluted in 110 µL extraction buffer. DNA extracts are stored at –20°C. (iii) Blood culture media and benzyl alcohol-guanidine hydrochloride organic extraction method132  A total of 0.1 mL of the inoculated medium is added to 0.1 mL of lysis buffer and is briefly mixed with a vortex mixer. Lysis buffer consists of 5.0 M guanidine hydrochloride—100 mM Tris (pH 8.0) in sterile water. A total of 0.4 mL of water is added, followed by the addition of 0.8 mL of 99% benzyl alcohol (Sigma), and the sample is mixed again by vortexing. The sample is centrifuged at 7000 × g for 5 min. A total of 0.4 mL of the aqueous supernatant is removed and is placed in a new microcentrifuge tube. A total of 40 µL of 3.0 M sodium acetate is added, followed by the addition of 440 µL of isopropanol, and the sample is centrifuged at 16,000 × g for 15 min at 4°C. The precipitated DNA is washed with 70% ethanol and the pellet is air-dried. The DNA is resuspended in 0.1 mL of 10 mM Tris–0.1 mM EDTA buffer at pH 8.5. 8.2.2.2  Detection Procedures 8.2.2.2.1  Identification of Cultured Strains Upon isolation of colonies, it is necessary to confirm the genus identity and also to further characterize to the species level. This characterization requires a battery of classical morphological and biochemical tests. The confidence level of the species identification will increase with the more tests that are carried out. Herein lies the greatest disadvantage of classical tools for identification of organisms, as even the most sophisticated array of tests can often lead to uncertainties in the classification of isolates.

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DNA-based techniques have as an advantage that the genomic material of an organism is investigated in a direct way, independent of culture conditions, making the process of identification more reliable. (i) Sequencing of 16S rRNA or cpn60 genes  DNA sequencing is the process of determining the order of the bases A, T, C, and G in a DNA strand. In essence, the DNA is used as a template to generate a set of fragments that differ in length from each other by a single base. The fragments are then separated by size and the bases at the end are identified, recreating the original sequence of the DNA. Adequate genes for the purpose of bacterial species identification are the 16S ribosomal RNA gene (rRNA) or chaperonin-60 (cpn60; also known as groEL or hsp60) genes. For identification, the sequence is compared to a public database (e.g., Genbank) or a curated database (e.g., SmartGene IDNS Bacteria) by using analysis software (BLAST148 or SmartGene, www. idns-smartgene.com). Sample Preparation and Applications for Detection of G. vaginalis. Although several universal primer pairs for amplification of the 16S ribosomal RNA gene have been published, not all of these primers are truly universal (reviewed by Srinivasan and Fredricks149 and Kalra150). For example, the universal primer 10f (5′-AGTTTGATCCTGGCTCAG-3′) allows amplification of G. vaginalis DNA from cultured bacteria, although it has three mismatches, but it fails to amplify this species from a complex mixture of bacteria due to competition with targets fully complementary to the primer.57 In our hands, the species-specific primer set GV10f (5′-GGTTCGATTCTGGCTCAG-3′)—534r (5′-ATTACCG CGGCTGCTGG-3′) provides the most efficient amplification of G. vaginalis.57,151 Amplification is performed in a 25  μL PCR mixture containing 0.1  μM of each primer, 12.5 μL of Promega master mix (Promega), 2.5 μL of DNA extract and distilled water. Thermal cycling consists of an initial denaturation of 5 min at 94°C, followed by 3 cycles of 1 min at 94°C, 2 min at 50°C, and 1 min at 72°C, followed by 35 cycles of 20 s at 94°C, 1 min at 50°C, and 1 min at 72°C, with a final extension of 10 min at 72°C and cooling to 10°C. PCR products are purified using the QIAquick PCR Purification Kit (Qiagen). Sequencing is done using the ABI Big Dye cycle sequencing reaction kit with AmpliTaq FS DNA polymerase (Applied Biosystems) with primer 534r. Sequencing reaction products are analyzed on an ABI 310 genetic analyzer (Applied Biosystems). Inspection of the electropherograms and comparison of the sequences to the 16S rRNA gene sequences in GenBank is done using SmartGene Software (http://www.smartgene.com/). (ii) tRNA-intergenic polymorphism length analysis (tDNA– PCR)  tDNA–PCR makes use of a single pair of universal primers—that is, primers complementary to conserved regions in the tRNA-genes of most bacterial groups—that are directed outwardly to amplify the spacers between the tRNA-genes. The tRNA-intergenic spacers are multiple

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per genome and have different lengths, such that amplification of the spacers from a bacterial genome results in a DNA-fingerprint consisting of multiple DNA-fragments immediately after amplification and electrophoresis.152 The obtained patterns are highly similar for the different strains within the same species, but differ between species. As such, amplification of tRNA-intergenic spacers, combined with high resolution automated electrophoresis, as is possible on (e.g., the ABI310 capillary electrophoresis system),153 makes it possible to construct a library with tDNA–PCR fingerprints of reference strains. Identification of unknown cultured strains can then be obtained by comparing the tDNA-fingerprint of the unknown with those present in the library.154,155 Special software has been developed to do so with a minimum of hands on time.156 We have developed such a library, covering most human bacterial species and several veterinary and environmental species. For most of the cultivable vaginal species, we have determined the corresponding tDNA–PCR patterns.98,157 Verhelst et al. and El Aila et al. applied tDNA–PCR identification for species cultured from the vaginal swabs of pregnant women for the purpose of microflora characterization.98,158 Vaginal swabs on Amies transport medium or swirled in liquid Amies transport medium (ESwab) were streaked onto rich agar plates and incubated anaerobically at 37°C upon arrival at the microbiology laboratory. After 4 days of incubation, all the isolates with different colony morphology were selected for identification. DNA was extracted by simple alkaline lysis (see crude DNA extraction). tDNA–PCR and capillary electrophoresis were carried out. This method was found to be a reproducible, cheap, and fast identification method for most vaginal species, including G. vaginalis. 8.2.2.2.2 Culture-Independent Detection and Identification (i) Conventional species-specific assays  In conventional PCR, detection and semiquantification of the amplified sequence are performed at the end of the reaction after the last PCR cycle and involve postPCR analysis, such as size fractionation of amplicons by gel electrophoresis and image analysis. Several assays for the detection of G. vaginalis, targeting respectively the 16S-23S rRNA spacer region58,60,159 or the 16S rRNA gene,160 were designed. Using species-specific PCR for G. vaginalis, targeting the 16S-23S rRNA spacer region, it could be shown that 40% of women, irrespective of their clinical status, and 10 out of 11 patients suffering from bacterial vaginosis, were carrying G. vaginalis.60 Through conventional PCR analysis of 515 vaginal samples, Verhelst et al.57,98 found that the presence of A. vaginae is a diagnostically more valuable marker for bacterial vaginosis than the presence of G.  vaginalis since respectively 14.7% of normal vaginal microflora sample and 86.4% of bacterial vaginosis samples showed an amplicon while the percentage of positive samples for G. vaginalis specific PCR in both categories were respectively 28.9% and 86.4%. For

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this purpose, G. vaginalis species-specific primers F-GV1 (5′-TTACTGGTGTATCACTGTAAGG-3′) and R-GV-3 (5′-CCGTCACAGGCTGAACAGT-3′) as designed by Zariffard et al.161 were used. Briefly, a 20 μL PCR mixture contained respectively 0.05 and 0.4  μM primers, 10 μL of Promega master mix (Promega), 2 μL of Qiagen DNA extract of the samples, and distilled water. Thermal cycling consisted of an initial denaturation of 10 min at 94°C, followed by 50 cycles of 5 s at 94°C, 45 s at 55°C, and 45 s at 72°C, and a final extension of 10 min at 72°C. Five μL of the amplified product was visualized on a 2% agarose gel. (ii) Real-time assays  Real-time polymerase chain reaction, also called quantitative polymerase chain reaction (qPCR), is a PCR procedure that enables both detection and quantification of a specific DNA target in a single tube, thereby limiting the risk of contamination and providing fast and reliable detection compared to classical end-point analysis PCR techniques. In contrast to conventional PCR, amplification is tracked during each replication cycle. Two common methods for detection of products in qPCR are (i) SYBRGreen, a fluorescent dye that intercalates with any double-stranded DNA, and (ii) sequence-specific DNA probes that are labeled with a fluorescent reporter that permits detection only after hybridization of the probe with its complementary DNA target. Using sequence-specific primers, the absolute number of chromosomes of a particular DNA target can be determined by extrapolating quantitative information obtained from a standard curve. Primer sets targeting the 16S rRNA gene or the 16S-23S spacer region of G. vaginalis were applied to assess the number of chromosomes of these microorganisms in DNA extracts obtained from vaginal swab specimens.61,159,161–165 De Backer et al.61 confirmed by qPCR, using the primers designed by Zariffard et al.,161 that the presence of A. vaginae is a diagnostically more valuable marker for bacterial vaginosis than the presence of G. vaginalis. The qPCR Core Kit for SYBR Green I (Eurogentec) was applied and analysis was performed on the ABI 7000 qPCR system (Applied Biosystems). PCR mixtures contained 2.5 μL of DNA extract, 2.5 μL of 10× Reaction Buffer, 3.5 mM MgCl2, 0.2  mM dNTP mixture, 0.625 U HotGoldStar Taq polymerase, 0.75 μL SYBR® Green I, diluted tenfold in DMSO and 0.2  µM primers, and were adjusted with HPLC grade water to 25 μL. Thermal cycling consisted of an initial denaturation of 10 min at 95°C, followed by 50 cycles of 45 s at 94°C, 45 s at 55°C, and 45 s at 72°C. Using the same reaction conditions, El Aila et al. (in preparation) assessed the presence of G. vaginalis in the vaginal and rectal microflora of 71 pregnant women and found that G. vaginalis was present in 74.6% of the women, either vaginally only (9.8%), rectal only (15.4%), or in both sites (49.2%), and concluded that these results confirm that the rectum serves as a reservoir for G. vaginalis and other bacterial vaginosis associated species. Interestingly, these authors found also a quantitative correspondence between rectum and vagina for this and other species.

Molecular Detection of Human Bacterial Pathogens

Other qPCR assays for quantitative assessment of G. vaginalis concentrations were published by Zariffard et al.,161 Trama et al.,162 Biagi et al.,163 Fredricks et al.,164 and ZozayaHinchliffe et al.166 Furthermore, the utility of qPCR as a diagnostic tool for bacterial vaginosis has been explored by Sha et al.,159 and Menard et al.165 Using 203 samples, from both of women with and without bacterial vaginosis, Sha et al. reported that increasing levels of G. vaginalis and M. hominis and decreasing levels of lactobacilli were significantly associated with bacterial vaginosis, with a sensitivity and specificity of 83% and 78% when compared with Nugent score. Menard et al.165 assessed the utility of G. vaginalis and A. vaginae loads as a diagnostic tool in 231 women and confirmed the results of De Backer et al.61 that the presence of A. vaginae at ≥108 copies/mL and G. vaginalis at ≥109 copies/ mL had a sensitivity and specificity of 95% and 99% when compared to Nugent scoring. (iii) Commercial point-of-care test for the diagnosis of G. vaginalis  The Affirm™ VP III Microbial Identification Test (BD Diagnostic Systems, NJ, USA) is a DNA hybridisation assay intended for detection and identification of Candida species, G. vaginalis and Trichomonas vaginalis in vaginal fluid specimens from patients with symptoms of bacterial vaginosis. The test uses two distinct single-stranded nucleic acid probes for each organism, a capture probe and a color development probe, that are complementary to unique genetic sequences of the target organisms. The capture probes are immobilized on beads embedded in a Probe Analysis Card (PAC), which contains a separate bead for each target organism. The color development probes are contained in a multiwell Reagent Cassette (RC). During sample preparation, the sample is treated with the Lysis Solution and heated. This process ruptures the walls of the organism, releasing the nucleic acid. A second solution, the Buffer Solution, is added. This solution stabilizes the nucleic acid and establishes the stringency conditions necessary for specific hybridization. Automated processing of the sample is performed using the BD MicroProbeTM Processor. The final step is reading the results of color development on each of the target organism beads and controls. The procedure requires between 30 and 45 min to complete and can be done in an office setting. This DNA hybridization test is positive only for concentrations of G. vaginalis in excess of 2 × 105 bacterial cells per ml of vaginal fluid167 and therefore should be positive most often in women with bacterial vaginosis and rarely in women with normal vaginal microflora. Promising results have been reported with this approach.167 These authors concluded that the probe could be used as a supplement to the Amsel and Nugent criteria for the diagnosis of bacterial vaginosis. (iv) Fluorescence in situ hybridization (FISH)  FISH can be used to visualize bacteria. Fluorescent probes are hybridized to mRNA or DNA present in cells and are visualized using epifluorescence microscopy. Data can be collected in both quantitative and qualitative modes. For example, when

Gardnerella

fluorescent probes are combined with flow cytometry, one can rapidly count and collect cells. With confocal scanning laser microscopy, one can visualize the spatial arrangement of cells in tissues or body fluids. FISH can also be used to study the bacterial community structure and the spatial organization of bacteria on the epithelial surfaces of vaginal biopsy specimens.31 Biopsy specimens are taken from the middle side wall of the vagina with biopsy forceps, fixed in nonaqueous Carnoy solution, and then processed and embedded into paraffin blocks. Four micrometer sections are placed on SuperFrost slides for FISH analysis. Oligonucleotide probes are synthesized with a carbocyanine dye (Cy3), fluorescein isothiocyanate, or Cy5 fluorescent dye at the 5′. The probe used for visualization of G. vaginalis (GardV) has the following sequence: 5′-CCA CCG TTA CAC CGC GAA-3. Hybridisation is performed using a washing temperature of 50°C and bacteria are detected by FISH and 4′,6-diamidino-2-phenylindole counter stain. This paper confirmed the strong link between A. vaginae, G. vaginalis, and bacterial vaginosis in a substantial manner, not only by showing the simultaneous presence of these two species in more than half of the cases of bacterial vaginosis, but also by showing that there was a spatial association in biofilms. Moreover, the realization that bacterial biofilms can be formed at the vaginal epithelium explains the presence of the well-known “clue cells,” designated as such because they are a hallmark for bacterial vaginosis. Finally, the insight that vaginosis is associated with bacterial biofilms may also nicely explain the recurrent character of vaginosis in many women. A more recent FISH study explained the clinical observation that, despite the temporary improvement of clinical symptoms of bacterial vaginosis by antibiotic treatment, the symptoms recur.106 It could be shown by FISH analysis of bacterial mRNA and DNA extracted from vaginal biopsies taken during treatment that the bacteria in the biofilm become dormant during treatment, but are not eradicated. After treatment, bacterial metabolic activity is regained and symptoms return. (v) Checkerboard DNA hybridization  A method allowing for simultaneous detection of multiple organisms is the checkerboard method.168 In this case, DNA of the complex microflora is extracted, spotted in separate lanes on a membrane, and hybridized with multiple labeled whole genomic DNA probes. A checkerboard for the simultaneous detection of G. vaginalis, P. bivia, B. ureolyticus, and M. curtisii was evaluated and it was concluded that hybridization was significantly associated with Gram-stain results since more species were detected by DNA hybridization as the Nugent score progressed from a normal to a disturbed microflora.169 Persson et al. assessed, by a semiquantitative DNA–DNA checkerboard hybridization assay of 74 bacterial species including G. vaginalis, whether the bacterial counts of oral and bacterial vaginosis species were higher in the vaginal microflora of women with gingivitis compared to healthy women. The load of G. vaginalis in the vaginal microflora of women with gingivitis was not elevated.170

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(vi) Oligonucleotide-coupled fluorescent microsphere  Hybridization can also take place on optically encoded microparticles.171 Pioneered by Luminex, these polystyrene microbeads are imbued with two fluorescent dyes and the ratio of these creates 100 unique spectral signatures. Each bead is coated with a different oligonucleotide probe. After PCR amplification of the targets, wherein amplicons are labeled, only those beads that have bound target will be detected by a flow cytometer. The main advantage of this approach is its ability to detect a large number of molecular targets simultaneously after a single PCR, making high-throughput analysis of multiple samples from large study groups or longitudinal studies technically feasible. A nine-plex Luminex assay was designed for characterizing the vaginal microflora and for exploring molecular diagnostics of bacterial vaginosis.172 DNA was extracted using Instagene Matrix (BioRad Laboratories). An amplicon for Luminex analysis was generated using two universal cpn60 primers sets. The G. vaginalis probe had the following sequence: 5′-CCAGCAACAACATTCTTTAAGCCTTCGTG-3′. The vaginal swab DNA extracts were used as templates for PCR amplification of the 5′ end of the cpn60 UT using two universal primers. Using the presence of one or both of A. vaginae and G. vaginalis as the defining criterion for bacterial vaginosis, these authors found that the method showed a high sensitivity and specificity for diagnosing bacterial vaginosis compared to traditional microscopy.

8.3  CONCLUSIONS AND FUTURE PERSPECTIVES While recent studies of the biofilm-forming potential and cytotoxic activity of G. vaginalis32 have renewed interest in the virulence potential of this organism, ongoing sequence analysis of the whole genome within the Human Microbiome Project and the G. vaginalis Genome Project (http://med. stanford.edu/sgtc/research/gardnerella_vaginalis.html) will further improve our understanding of the pathogenic role of G. vaginalis in bacterial vaginosis.173 This sensitive highthroughput analysis through deep pyrosequencing will further enlighten our knowledge of bacterial diversity and ecological dynamics associated with reproductive tract infection. Knowledge obtained by this metagenomic approach might then be translated into rapid diagnostic assays for profiling a selected set of microbial communities in a lab-based setting. To further explore the role of G. vaginalis in bacterial vaginosis, microarrays like the TNO vaginal microarray (V-chip, TNO, Zeist, Netherlands) offer the possibility to detect over 400 vaginal species. In conclusion, molecular diagnosis of bacterial vaginosis, with a central role for G. vaginalis and A. vaginae (see Chapter 4) is now pending.61,62,161,165,172 The study by Menard et al. that assessed the utility of quantitation of G. vaginalis and A. vaginae as a diagnostic tool for bacterial vaginosis clearly seems to provide a sound basis.149 However, as has been recently reiterated by Kalra et al., distinct bacterial

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vaginosis profiles consisting of differing species might predispose to different health outcomes.149 In cases of bacteremia, molecular detection methods should become the golden standard to detect and identify G. vaginalis, as blood culture media often contain inhibitors that prevent isolation of this species. Importantly, when changing traditional methods for molecular detection methods, special attention should be addressed to proper nucleic acid extraction methods.

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92 82. Lagace-Wiens, P.R. et al., Gardnerella vaginalis bacteremia in a previously healthy man: Case report and characterization of the isolate, J. Clin. Microbiol., 46, 804, 2008. 83. Hodge, T.W.J., Levy, C.S., and Smith, M.A., Disk space infection due to Gardnerella vaginalis, Clin. Infect. Dis., 21, 443, 1995. 84. Graham, S. et al., Vertebral osteomyelitis and discitis due to Gardnerella vaginalis, J. Med. Microbiol., 58, 1382, 2009. 85. Sivadon-Tardy, V. et al., Gardnerella vaginalis acute hip arthritis in a renal transplant recipient, J. Clin. Microbiol., 47, 264, 2009. 86. Francois, S., Guyadier-Souguieres, G., and Marcelli, C., Reactive arthritis due to Gardnerella vaginalis. A case-report, Rev. Rhum. Engl. Ed., 64, 138, 1997. 87. Schapira, D., Braun-Moscovici, Y., and Nahir, A.M., Reactive arthritis induced by Gardnerella vaginalis, Clin. Exp. Rheumatol., 20, 732, 2002. 88. Toussirot, E., Plesiat, P., and Wendling, D., Reiter’s syndrome induced by Gardnerella vaginalis, Scand. J. Rheumatol., 27, 316, 1998. 89. Verstraelen, H. et al., The epidemiology of bacterial vaginosis in relation to sexual behaviour, BMC Microbiol., 10, 81, 2010. 90. Swidsinski A, et al. Gardnerella biofilm involves females and males and is transmitted sexually. Gynecol Obstet Invest., 70, 256, 2010. ‌ 91. Bartley, D.L., Morgan, L., and Rimsza, M.E., Gardnerella vaginalis in prepubertal girls, Am. J. Dis. Child., 141, 1014, 1987. 92. Gardner, J.J., Comparison of the vaginal flora in sexually abused and nonabused girls, J. Pediatr., 120, 872, 1992. 93. Steele, A.M., and de San Lazaro, C., Transhymenal cultures for sexually transmissible organisms, Arch. Dis. Child., 71, 423, 1994. 94. Ingram, D.L. et al., Gardnerella vaginalis infection and sexual contact in female children, Child Abuse Negl., 16, 847, 1992. 95. Shafer, M.A. et al., Microbiology of the lower genital tract in postmenarchal adolescent girls: Differences by sexual activity, contraception, and presence of nonspecific vaginitis, J. Pediatr., 107, 974, 1985. 96. Bump, R.C., Sachs, L.A., and Buesching, W.Jr., Sexually transmissible infectious agents in sexually active and virginal asymptomatic adolescent girls, Pediatrics, 77, 488, 1986. 97. Yen, S. et al., Bacterial vaginosis in sexually experienced and non-sexually experienced young women entering the military, Obstet. Gynecol., 102, 927, 2003. 98. Verhelst, R. et al., Comparison between Gram stain and culture for the characterization of vaginal microflora: Definition of a distinct grade that resembles grade I microflora and revised categorization of grade I microflora, BMC Microbiol., 5, 61, 2005. 99. Fredricks, D.N., Fiedler, T.L., and Marrazzo, J.M., Molecular identification of bacteria associated with bacterial vaginosis, N. Engl. J. Med., 353, 1899, 2005. 100. McCaffrey, M. et al., Bacterial vaginosis in lesbians: evidence for lack of sexual transmission, Int. J. STD AIDS, 10, 305, 1999. 101. Marrazzo, J.M. et al., haracterization of vaginal flora and bacterial vaginosis in women who have sex with women, J. Infect. Dis., 185, 1307, 2002. 102. Evans, A.L. et al., Prevalence of bacterial vaginosis in lesbians and heterosexual women in a community setting, Sex. Transm. Infect., 83, 470, 2007.

Molecular Detection of Human Bacterial Pathogens 103. Weyers, S. et al., Microflora of the penile skin-lined neova­ gina of transsexual women, BMC Microbiol., 9, 102, 2009. 104. Dawson, S.G. et al., Male carriage of Gardnerella vaginalis, Br. J. Vener. Dis., 58, 243, 1982. 105. Holst, E., Goffeng, A.R., and Andersch, B., Bacterial vaginosis and vaginal microorganisms in idiopathic premature labor and association with pregnancy outcome, J. Clin. Microbiol., 32, 176, 1994. 106. Swidsinski, A. et al., An adherent Gardnerella vaginalis biofilm persists on the vaginal epithelium after standard therapy with oral metronidazole, Am. J. Obstet. Gynecol., 198, 97.e1, 2008. 107. Schwebke, J.R., Rivers, C., and Lee, J., Prevalence of Gardnerella vaginalis in male sexual partners of women with and without bacterial vaginosis, Sex. Transm. Dis., 36, 92, 2009. 108. Chattopadhyay, B., and Teli, J.C., Isolation of Gardnerella vaginalis from routine genito-urinary tract specimens, J. Infect., 8, 157, 1984. 109. Ison, C.A., and Easmon, C.S., Carriage of Gardnerella vaginalis and anaerobes in semen, Genitourin Med., 61, 120, 1985. 110. Elsner, P., and Hartmann, A.A., Gardnerella vaginalis in the male upper genital tract: A possible source of reinfection of the female partner, Sex. Transm. Dis., 14, 122, 1987. 111. Lam, M.H., Birch, D.F., and Fairley, K.F., Prevalence of Gardnerella vaginalis in the urinary tract, J. Clin. Microbiol., 26, 1130, 1988. 112. Hillier, S.L. et al., Relationship of bacteriologic characteristics to semen indices in men attending an infertility clinic, Obstet. Gynecol., 75, 800, 1990. 113. Kjaergaard, N. et al., Microbiology of semen specimens from males attending a fertility clinic, APMIS, 105, 566, 1997. 114. Andrade-Rocha, F.T., Colonization of Gardnerella vaginalis in semen of infertile men: Prevalence, influence on sperm characteristics, relationship with leukocyte concentration and clinical significance, Gynecol. Obstet. Invest., 68, 134, 2009. 115. Virecoulon, F. et al., Bacterial flora of the low male genital tract in patients consulting for infertility, Andrologia, 37, 160, 2005. 116. Schwebke, J.R., New concepts in the etiology of bacterial vaginosis, Curr. Infect. Dis. Rep., 11, 143, 2009. 117. Verstraelen, H., Bacterial vaginosis: a sexually enhanced disease, Int. J. STD AIDS, 19, 575, 2008. 118. Salmon, S.A. et al., Isolation of Gardnerella vaginalis from the reproductive tract of four mares, J. Vet. Diagn. Invest., 2, 167, 1990. 119. Golberg, R.L., and Washington, J.A.I.I., Comparison of isolation of Haemophilus vaginalis (Corynebacterium vaginale) from peptone-starch-dextrose agar and Columbia colistin-nalidoxic acid agar, J. Clin. Microbiol., 4, 245, 1976. 120. Piot, P. et al., Identification of Gardnerella (Haemophilus) vaginalis, J. Clin. Microbiol., 15, 19, 1982. 121. Totten, P.A. et al., Selective differential human blood bilayer media for isolation of Gardnerella (Haemophilus) vaginalis, J. Clin. Microbiol., 15, 141, 1982. 122. Piot, P., and Van Dyck, E., Isolation and identification of Gardnerella vaginalis, Scand. J. Infect. Dis., 40, S15, 1983. 123. Holst, E., Reservoir of four organisms associated with bacterial vaginosis suggests lack of sexual transmission, J. Clin. Microbiol., 28, 2035, 1990. 124. Ison, C.A. et al., Comparison of culture and microscopy in the diagnosis of Gardnerella vaginalis infection, J. Clin. Pathol., 35, 550, 1982.

Gardnerella 125. Taylor, E., and Phillips, I., The identification of Gardnerella vaginalis, J. Med. Microbiol., 16, 83, 1983. 126. Bailey, R.K., Voss, J.L., and Smith, R.F., Factors affecting isolation and identification of Haemophilus vaginalis (Corynebacterium vaginale), J. Clin. Microbiol., 9, 65, 1979. 127. Gibbs, R.S. et al., Microbiologic and serologic studies of Gardnerella vaginalis in intra-amniotic infection, Obstet. Gynecol., 70, 187, 1987. 128. La Scolea, L.J.J., Dryja, D.M., and Dillon, W.P., Recovery of Gardnerella vaginalis from blood by the quantitative direct plating method, J. Clin. Microbiol., 20, 568, 1984. 129. Legrand, J. C. et al., Gardnerella vaginalis bacteremia from pulmonary abscess in a male alcohol abuser, J. Clin. Microbiol., 27, 1132, 1989. 130. Reimer, L.G., and Reller, L.B., Gardnerella vaginalis bacteremia: A review of thirty cases, Obstet. Gynecol., 64, 170, 1984. 131. Reimer, L.G., and Reller, L.B., Effect of sodium polyanetholesulfonate and gelatin on the recovery of Gardnerella vaginalis from blood culture media, J. Clin. Microbiol., 21, 686, 1985. 132. Fredricks, D.N., and Relman, D.A., Improved amplification of microbial DNA from blood cultures by removal of the PCR inhibitor sodium polyanetholesulfonate, J. Clin. Microbiol., 36, 2810, 1998. 133. Boskey, E.R. et al., Acceptability of a self-sampling technique to collect vaginal smears for gram stain diagnosis of bacterial vaginosis, Womens Health Issues, 14, 14, 2004. 134. Jones, H.E. et al., Home-based versus clinic-based selfsampling and testing for sexually transmitted infections in Gugulethu, South Africa: randomised controlled trial, Sex. Transm. Infect., 83, 552, 2007. 135. Baay, M.F. et al., Feasibility of collecting self-sampled vaginal swabs by mail: Quantity and quality of genomic DNA, Eur. J. Clin. Microbiol. Infect. Dis., 28, 1285, 2009. 136. Forney, L.J. et al., Comparison of self-collected and physician-collected vaginal swabs for microbiome analysis, J. Clin. Microbiol., 2010. 137. El Aila, N.A. et al., Genotyping of Streptococcus agalactiae (group B streptococci) isolated from vaginal and rectal swabs of women at 35–37 weeks of pregnancy, BMC Infect. Dis., 9, 153, 2009. 138. Van Dyck, E. et al., Evaluation of the Roche Neisseria gonorrhoeae 16S rRNA PCR for confirmation of AMPLICOR PCR-positive samples and comparison of its diagnostic performance according to storage conditions and preparation of endocervical specimens, J. Clin. Microbiol., 39, 2280, 2001. 139. Van Dyck, E. et al., Detection of Chlamydia trachomatis and Neisseria gonorrhoeae by enzyme immunoassay, culture, and three nucleic acid amplification tests, J. Clin. Microbiol., 39, 1751, 2001. 140. de Boer, R. et al., Improved detection of microbial DNA after bead-beating before DNA isolation, J. Microbiol. Methods, 80, 209, 2010. 141. Fredricks, D.N., Smith, C., and Meier, A., Comparison of six DNA extraction methods for recovery of fungal DNA as assessed by quantitative PCR, J. Clin. Microbiol., 43, 5122, 2005. 142. Fliss, I. et al., A rapid and efficient method of lysis of Listeria and other gram-positive bacteria using mutanolysin, Biotechniques, 11, 453, 456, 1991. 143. Akane, A. et al., Identification of the heme compound copurified with deoxyribonucleic acid (DNA) from bloodstains, a major inhibitor of polymerase chain reaction (PCR) amplification, J. Forensic Sci., 39, 362, 1994.

93 144. Wiedbrauk, D.L., Werner, J.C., and Drevon, A.M., Inhibition of PCR by aqueous and vitreous fluids, J. Clin. Microbiol., 33, 2643, 1995. 145. Holodniy, M. et al., Inhibition of human immunodeficiency virus gene amplification by heparin, J. Clin. Microbiol., 29, 676, 1991. 146. Khan, G. et al., Inhibitory effects of urine on the polymerase chain reaction for cytomegalovirus DNA, J. Clin. Pathol., 44, 360, 1991. 147. Ahokas, H., and Erkkila, M.J., Interference of PCR amplification by the polyamines, spermine and spermidine, PCR Methods Appl., 3, 65, 1993. 148. Altschul, S.F. et al., Basic local alignment search tool, J. Mol. Biol., 215, 403, 1990. 149. Srinivasan, S., and Fredricks, D.N., The human vaginal bacterial biota and bacterial vaginosis, Interdiscip. Perspect. Infect. Dis., 2008, 750479, 2008. 150. Kalra, A. et al., Bacterial vaginosis: Culture- and PCRbased characterizations of a complex polymicrobial disease’s pathobiology, Curr. Infect. Dis. Rep., 9, 485, 2007. 151. Verstraelen, H. et al., Longitudinal analysis of the vaginal microflora in pregnancy suggests that L. crispatus promotes the stability of the normal vaginal microflora and that L. gasseri and/or L. iners are more conducive to the occurrence of abnormal vaginal microflora, BMC Microbiol., 9, 116, 2009. 152. Welsh, J., and McClelland, M., Genomic fingerprints produced by PCR with consensus tRNA gene primers, Nucleic Acids Res., 19, 861, 1991. 153. Vaneechoutte, M. et al., Comparison of PCR-based DNA fingerprinting techniques for the identification of Listeria species and their use for atypical Listeria isolates, Int. J. Syst. Bacteriol., 48, 127, 1998. 154. Baele, M. et al., Application and evaluation of the interlaboratory reproducibility of tRNA intergenic length polymorphism analysis (tDNA-PCR) for identification of Streptococcus species, J. Clin. Microbiol., 39, 1436, 2001. 155. Baele, M. et al., tDNA-PCR followed by automated fluorescent capillary electrophoresis for identification of bacterial species, CSH Protoc., 4, prot5196, 2009. 156. Baele, M. et al., Application of tRNA intergenic spacer PCR for identification of Enterococcus species, J. Clin. Microbiol., 38, 4201, 2000. 157. Baele, M. et al., Identification of Lactobacillus species using tDNA-PCR, J. Microbiol. Methods, 50, 263, 2002. 158. El Aila, N.A. et al., Identification and genotyping of bacteria from paired vaginal and rectal samples from pregnant women indicates similarity between vaginal and rectal microflora, BMC Infect. Dis., 9, 167, 2009. 159. Sha, B.E. et al., Utility of Amsel criteria, Nugent score, and quantitative PCR for Gardnerella vaginalis, Mycoplasma hominis, and Lactobacillus spp. for diagnosis of bacterial vaginosis in human immunodeficiency virus-infected women, J. Clin. Microbiol., 43, 4607, 2005. 160. Nath, K., Sarosy, J.W., and Stylianou, S.P., Suitability of a unique 16S rRNA gene PCR product as an indicator of Gardnerella vaginalis, Biotechniques, 28, 222, 2000. 161. Zariffard, M.R. et al., Detection of bacterial vaginosis-related organisms by real-time PCR for Lactobacilli, Gardnerella vaginalis and Mycoplasma hominis, FEMS Immunol. Med. Microbiol., 34, 277, 2002. 162. Trama, J.P. et al., Rapid detection of Atopobium vaginae and association with organisms implicated in bacterial vaginosis, Mol. Cell. Probes, 22, 96, 2008.

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9 Gordonia Brent A. Lasker, Benjamin D. Moser, and June Brown CONTENTS 9.1 Introduction........................................................................................................................................................................ 95 9.1.1 Classification........................................................................................................................................................... 95 9.1.2 Morphology............................................................................................................................................................ 96 9.1.3 Chemotaxonomic Characteristics........................................................................................................................... 96 9.1.4 Epidemiology.......................................................................................................................................................... 96 9.1.5 Diagnosis................................................................................................................................................................ 98 9.1.5.1 Phenotypic, Biochemical, and Susceptibility Studies.............................................................................. 98 9.1.5.2 Molecular Techniques for Identification and Phylogenetic Analysis..................................................... 102 9.2 Methods............................................................................................................................................................................ 105 9.2.1 Sample Preparation............................................................................................................................................... 105 9.2.2 Detection Procedures............................................................................................................................................ 105 9.3 Conclusions and Future Perspectives............................................................................................................................... 107 References.................................................................................................................................................................................. 108

9.1  INTRODUCTION The genus Gordonia was originally described in 1971 by Tsukamura following the isolation of organisms from sputa of patients with pulmonary disease and from soil.1 The name of the genus was chosen to honor Ruth E. Gordon, an American microbiologist. There are currently 28 validly described species in this genus, and Gordonia bronchialis is the type species. According to the phylogenetic position of the genus based on the analysis of 16S rRNA sequences in 1997 by Stackebrandt et al., Gordonia is the type genus of the family Gordoniaceae within the suborder Corynebacterineae and of the order Actinomycetales.2 In this same publication, these authors corrected the original spelling to Gordonia.2 Most species are not yet associated with human infection because 15 of the 17 species identified in the last 10 years have been isolated from environmental sources. The clinically significant species are the primary focus of this chapter but the environmental species are included in comparative biochemical and susceptibility tables. An increased recognition of Gordonia species may result in delineation of species-specific differences, geographical distribution, disease associations, and greater awareness of antimicrobial susceptibility profiles. The biology of this genus has recently been reviewed with an emphasis on its potential usefulness in environmental and industrial biotechnology.3

9.1.1  Classification In 1971, as originally described and named, the Gordona genus  was composed of Gordona bronchialis, G. rubra

(Gordonia rubripertincta), and G. terrae.1 Tsukamura ­separated these species from rapidly growing mycobacteria by their slight acid fastness and their inability to produce arylsulfatase. In 1977, Goodfellow and Alderson transferred these members of Gordona, Mycobacterium rhodochrous, and the “rhodochrous” complex to the genus Rhodococcus based on extensive numerical computerized analysis of 92 characteristic traits.4 The three species, originally assigned to the genus Gordona, were studied by Collins et al.5 The study revealed that these species contained mycolic acids with a range from 48 to 66 carbon atoms and major amounts of dehydrogenated menaquinones with nine isoprene units [MK-9 (H2)], therefore leaving the remaining species of the genus Rhodococcus more homogeneous, with their mycolic acids ranging from 34 to 52 carbon atoms and with a major menaquinone of only eight isoprene units [MK-8 (H2)]. The mycolic acid and menaquinone analyses were in agreement with the data derived from the 16S rRNA gene sequencing of Stackebrandt et al. and led to the revival of the genus Gordona Tsukamura 1971.6 R. bronchialis, R. rubra, and R. terrae were reclassified to the genus Gordona and, additionally, Rhodococcus sputi was reclassified as Gordona sputi.6,7 The transferring of Nocardia amarae Lechevalier and Lechevalier 1974 and Rhodococcus aichiensis Tsukamura 1982 to the genus Gordona by Klatte et al. in 1994, based on 16S rRNA gene sequencing analysis and chemotaxonomic methods, further supported the integrity of the genus Gordona.8–10 Two additional species added to the genus Gordona were Gordona hydrophobica11 and G. hirsuta.12 The correct epithet was introduced in 1997 as the genus Gordonia.2 New species increased steadily from 1997 95

96

to 2009: Gordonia rhizophera,13 Gordonia alkanivorans,14 Gordonia desulfuricans,15 Gordonia polyisoprenivorans,16 Gordonia amicalis,17 Gordonia namibiensis,18 Gordonia sihwensis,19 Gordonia sinesedis,20 Gordonia westphalia,21 Gordonia paraffinivorans,22 Gordonia otitidis,23 Gordonia araii,24 Gordonia defluvii,25 Gordonia effusa,24 Gordonia soli,26 Gordonia shandongensis,27 Gordonia malaquae,28 Gordonia lacunae,29 Gordonia cholesterolivorans,30 and Gordonia kroppenstedtii.31 The taxonomic and clinical laboratory identification of the genus Gordonia today is dependent on phenotypic, chemotaxonomic, and molecular characterization.

9.1.2  Morphology The microscopic and colonial morphology of the individual species of Gordonia is very similar to that of Rhodococcus. Colonial morphologies observed either microscopically or macroscopically are species dependent and range from slimy to smooth or glossy and may vary from smooth to rough after 2–3 days incubation on a variety of media (e.g., blood agar media, Middlebrook 7H11 agar, brain heart infusion and heart infusion agar). The color of the colonies, often not apparent until at least 7 days of incubation at optimal temperature, also varies through a wide range, from white, cream, beige, or pink, to the more classic orange-salmon color of most Gordonia species. The organisms are gram-positive with coccoid to bacillary forms that may be weakly acid fast (4–10 organisms per oil immersion field) with the modified Kinyoun (1% sulfuric acid) stain. Briefly, all members of the genus are aerobic, nonsporeforming, gram-positive nocardioforms with an oxidative carbohydrate metabolism.

9.1.3  Chemotaxonomic Characteristics In addition to phenotypic features, there are a number of salient properties that are important in determining genus assignment. The chemotaxonomic characteristics that separate the genus Gordonia from related genera have been reviewed by Arenkötter et al.3 and Takeuchi and Hatano.13 Important differential traits include whole cell wall composition chemotype IV Lechevalier and Lechevalier32 (contains arabinose and galactose as major sugars and meso-diaminopimelic acid as the only diamino acid); the presence of MK-9 (H2) as the predominant menaquinone; saturated, unsaturated, and tuberculostearic acid in whole-cell fatty acid composition; mycolic acids with a range of 48–66 carbon atoms; N-glycolated muramic acid; presence of phosphatidylethanolamine; and G + C content of DNA ranging from 63 to 69 mol%.

9.1.4  Epidemiology Members of the genus Gordonia were originally regarded as opportunistic pathogens and were isolated from sputum of humans with respiratory diseases (e. g., G. bronchialis, G. aichiensis, and G. sputi). Recently, more species have been isolated with the ability to degrade xenobiotics,

Molecular Detection of Human Bacterial Pathogens

environmental pollutants or slowly biodegradable natural polymers (e.g., benzothiophene and dibenzothiophene by G.  desulfuricans and G. amicalis, alkanes by G. alkanivorans, malodorous animal emissions such as aldehydes and ketones by G. hydrophobica and G. hirsuta, polyisoprenoids and synthetic cis-1,4-polyisoprene by G. polyisoprenivorans and G.  westfalica, nitrile compounds by G. namibiensis, hydrocarbons such as liquid paraffin by G. paraffinivorans,22 and phenol compounds by G. kroppenstedtii).31 Given the difficulties of accurately identifying Gordonia species by phenotypic tests alone, we may not be certain of the identification made by earlier investigators of Gordonia clinical isolates, especially those made without molecular studies. However, we present and review case reports that claim an association of certain gordoniae with clinical disease. Gordonia araii. The type strain for this species, G. araii IFM 10211T,24 was first isolated from a sputum specimen in a 48-year-old male Japanese patient with bacterial pneumonia. The specimen underwent extensive polyphasic taxonomic analysis and was identified as a novel species based on 16S rRNA gene sequence analysis and DNA–DNA hybridization.24 Pathogenic G. araii also appears in the literature as the disease causative agent in a 27-year-old following a sport-related injury of the right anterior cruciate ligament (ACL) and right medial meniscus tear.33 After surgery, the patient complained of fever and knee pain, which was followed 10 weeks later by erythema and edema of the right knee. At 21 weeks after surgery, MRI revealed an effusion and edema in both the tibia and femur of the graft attachment site. All foreign materials related to the surgical procedure were removed and cultured; growth of a gram-positive bacillus was observed on brain heart infusion agar with 5% sheep blood after 3 days. This isolate was resistant to azithromycin (>256.0 ug/mL) and was treated with intravenous trimethoprim/sulfamethoxazole (175–875 mg twice daily). This was the first case of confirmed G. araii infection associated with a medical device in an immunocompetent individual.33 Gordonia aichiensis. Gordonia aichiensis (type strain E9028T) was first isolated from human sputum and initially identified as Rhodococcus aichiense.10 The transfer from Rhodococcus to the genus Gordona was described by Klatte et al.8 Clinical presentation of G. aichiensis is rare in the literature. Two recorded cases are of a 74-year-old female and a 72-year-old male with G. aichiensis cultured from sputum and pus, respectively.34 Gordonia bronchialis. Gordona (Gordonia) bronchialis was first proposed as a new species by Tsukamura in 1971 after a number of organisms, isolated from patients with pulmonary disease, grew with intermediate characteristics between the genera Mycobacterium and Nocardia.1 Gordonia bronchialis was identified as a human diseasecausing agent in sternal wound infections following open heart surgery.35 In these seven cases, Richet et al. determined that a noninfected circulating nurse and her dogs were the reservoirs for the organism. The nurse was inadvertently transmitting the infectious organism by touching a colonized area of her body, such as her scalp, and then

Gordonia

accidentally transferred the organism to the water bath used in a standard clotting test during open heart surgery procedures and contaminated instruments and materials used on the patient.35 In 2004, G. bronchialis was cultured from the blood of a 58-year-old woman with recently diagnosed diabetes mellitus presenting with breathlessness, fever, and chills related to a sequestrated lung.36 Known to be pathogenic in immunocompromised individuals,37 G. bronchialis has also been reported as the infectious agent in an immunocompetent individual with a recurrent breast abscess38 and in a 45-day-old premature neonate with apnea and bradycardia.39 Aoyama et al. characterized 31 Gordonia species clinical isolates from 1998 to 2008.34 Of the isolated strains, 13 were identified as G. bronchialis and were isolated from sites such as pleural fluid, sputum, granuloma, blood, skin, and pus. This represents the largest series of G. bronchialis clinical infections. Gordonia effusa. The type strain for G. effusa IFM 10200T24 was first isolated from a sputum specimen from a 74-year-old male Japanese patient with kidney dysfunction. Described concurrently with G. araii, the initial isolated specimen underwent extensive polyphasic taxonomic analysis and was identified as a novel species based on 16S rRNA gene sequence analysis and DNA–DNA hybridization.24 Gordonia effusa infection is rare and presents only this one time throughout the literature. Gordonia otitidis. Gordonia otitidis is an uncommon clinical infection. The type strain of G. otitidis, IFM 10032T, was first isolated from ear discharge in a 28-year-old Japanese female with external otitis in 2000.23 Also, G. otitidis was isolated from pleural fluid in a 60-year-old Japanese male with bronchitis, assumed to be the same case identified by Aoyama et al. in their investigation of G. otitidis found in pleural fluid during their study of 31 Gordonia clinical isolates.24 As recent as 2007, Blaschke et al. presented a case of infection in an 11-year-old who presented febrile with a history of central venous catheter use.39 Gordonia polyisoprenivorans. Gordonia polyisoprenivorans (type strain Kd2T) was first isolated from fouling water inside of an automobile tire.16 Several other rubberdegrading Gordonia species have been previously identified in automobile tires and from the soil of a rubber tree plantation in Vietnam.40,41 Although originally isolated from the environment, G. polyisoprenivorans appears in the literature as the disease-causing agent in clinical infection. Kempf et al. reported the first case of septicemia caused by G. polyisoprenivorans following a bone marrow transplant in a 26-year-old female.42 The patient was suffering from chronic myelogenous leukemia and received an allogenic bone marrow transplantation in December 2001. Before transplantation, a Hickman catheter was inserted into the patient and used for a myeloablative condition regimen. In March 2002, after presenting with a fever, blood samples were cultured from her Hickman catheter revealing G. polyisoprenivorans.42 Verma et al. describes an immunocompromised 78-year-old male who underwent surgery for an incarcerated left inguinal hernia.43 The patient had a right

97

internal jugular Hickman catheter inserted for blood transfusions during his procedure. Later, this patient underwent evaluation of recurrent anemia and a third-degree heart block. Echocardiography revealed vegetations on the aortic and mitral valve for which the patient underwent aortic valve replacement and debridement of the mitral valve, respectively. Histopathological examination showed acute endocarditis and blood culture grew a beaded gram-positive bacillus; later identified at Centers for Disease Control and Prevention (CDC) as G. polyisoprenivorans. Most recently, Gupta et al. presented the first case of pneumonia with associated bacteremia caused by G. polyisoprenvirans following long-term use of a central catheter.44 Gordonia rubripertincta. Gordonia rubripertincta (type strain ATCC 14352) was first described in clinical infection by Hart et al. reporting a case of a lung infection resembling tuberculosis caused by Rhodococcus rubropertinctus.45 The patient showed no immunosuppression, which is unusual for disease caused by the “rhodochrous” complex.45 The organism was identified as the primary pathogen in the abortion of 9-month-old fetus in a 13-year-old quarter horse. Samples of the lung following necropsy yielded the growth of a grampositive, club-shaped branching organism.46 Rhodococcus rubropertinctus was transferred into the genus Gordona in 1989 and renamed Gordonia in 1997.2 Gordonia sputi. Gordonia sputi is one of the most ­common disease-causing agents among the gordoniae. First described by Tsukamura in 1978 as Rhodococcus sputi, this organism has caused a series of human and animal ­infections. Tsukamura et al. describe a case of mesenteric lymphadenitis in a 6-month-old swine caused by R. sputi.47 Riegel et al. reported on the first case of bacteremia due to a strain of Gordona sputi in an immunodeficient patient.48 Riegel et al. describe a 34-year-old man with ocular melanoma and hepatic metastases, who was undergoing treatment prior to infection. Following 4 months of treatment with cis-platinum and intravenous interleukin-2, the patient developed a 40°C fever; blood culture grew a gram-positive coryneform bacterium. One week later, after another fever, three blood cultures yielded the same organism identified as G. sputi.48 Kuwabara et al. described a case of mediastinitis caused by G. sputi infection in a 54-year-old man subsequent to coronary artery bypass grafting49; followed by a Lesens et al. report of the first case of endocarditis.50 Lesens et al. described a 31-year-old woman with genetic blood disorders, hepatitis C infection, and hemochromatosis ­complicated by diabetes, adrenal insufficiency, and peripheral ­neuropathy presenting with mitral valve vegetations following the implantation of a central venous port. The site of the port showed no signs of infection; however, blood cultures from her central catheter and a peripheral source grew ­gram-­positive coryneform bacterium (G. sputi). More recently, cases of G. sputi bacteremia were reported in Brust et al.,37 as well as in 14 of the 31 clinical strains investigated by Aoyama et al.34 G. sputi again appears in a series of infections caused by Gordonia species in a Taiwan medical center.51 Of the 66 clinical isolates previously identified

98

as  Rhodococcus ­species, 15  were identified as Gordonia species; eight of them were G. sputi.51 Gordonia terrae. Gordonia terrae is a ubiquitous disease agent, causing brain abscesses,52 central nervous symptoms,53 and bacteremia 54–56 in humans. Drancourt et al. presented the case of an immunocompromised boy undergoing antineoplastic chemotherapy for a cerebral rhabdoid tumor that was complicated by the formation of a brain abscess caused by G. terrae infection.52 A 40-year-old immunocompetent woman had G. terrae isolated from central spinal fluid (CSF) in her evaluation of hemiparesis and brain abscesses that proved difficult to treat.53 A rare cause of mycetoma occurred in a 15-year-old boy in Sierra Leone following an injury to his left thumb from a thorn,57 and an even rarer example of G. terrae infection occurred in 2009 following the nipple piercing of a 29-year-old female.58 The granulomatous mastitis caused by the nipple piercing is the first reported Gordonia species-associated mastitis in the literature.58 Aoyama et al. described G. terrae infection in 1 of the 31 investigated samples in their characterization study,34 Lai et al. described G. terrae as the organism responsible for 7 of the 15 clinical Gordonia isolates in a Taiwan medical center,51 and Blaschke et al. presented four episodes of G. terrae bacteremia in three children, all with central venous catheters.39 The recurrence of G. terrae bacteremia in one child suggests that therapy may need to be prolonged compared to other organisms.

9.1.5  Diagnosis 9.1.5.1  P  henotypic, Biochemical, and Susceptibility Studies With an increasing number of Gordonia species and related genera, the use of only a limited number of phenotypic traits to obtain exact species or genus level identification has become impossible. However, the production of aerial hyphae on solid media and the production of branched filaments by members of the genus Nocardia are useful in differentiating them from the gordoniae (except for G. defluvii, which has both aerial hyphae and branched filaments). Distinguishing Gordonia species from related genera Corynebacterium, Mycobacterium, Rhodococcus, and Tsukamurella is more problematic. Gordonia can be separated from Mycobacterium and Tsukamurella as Gordonia are not 100% acid fast with modified Kinyoun. In addition, Gordonia strains do not produce arylsulfatase, as do Mycobacterium strains. Gordonia species can be distinguished from Corynebacterium species by a combination of chemotaxonomic (Gordonia species have mycolic acids with 48–66 carbon atoms while Corynebacterium species contain 22–36 carbon atoms) and morphological features (Gordonia species usually have a rod-to-coccoid cycle while Corynebacterium species have club-shaped, palisade, or angular arrangements). The microscopic and colonial morphology of the individual species of Gordonia is very similar to that of Rhodococcus emphasizing the difficulty in differentiating gordoniae from the rhodococci

Molecular Detection of Human Bacterial Pathogens

without chemotaxonomic (menaquinone and HPLC analysis) and molecular methods. Traditionally, the gordoniae were identified to the species level using a battery of biochemical tests including hydrolysis of adenine, casein, hypoxanthine, tyrosine, and xanthine.59 The two major types of carbohydrate utilization tests used to distinguish among the species of Gordonia included those of Gordon (assimilation and assimilation with acid production) and those of Goodfellow (assimilation only).60 However, because of the generally nonreactive nature of most species to these tests and because of the increasing number of described species, species assignment using biochemical methods alone does not provide reliable identification of all the currently recognized species. (See Table 9.1 for useful tests to establish a tentative identification of the gordoniae including the major clinically relevant species.) The Clinical Laboratory Standards Institute (CLSI, formerly NCCLS) has published an approved standard for susceptibility testing of mycobacteria, nocardiae, and other aerobic actinomycetes.61 The recommended method for gordoniae is broth microdilution.61 Antimicrobial agents recommended for testing include amikacin, amoxicillinclavulanate, ceftriaxone, ciprofloxacin, clarithromycin, imipenem, linezolid, minocycline, and trimethoprim/sulfamethoxazole. Interpretive breakpoints of these agents are provided and a recommended quality control procedure is specified.61 The antimicrobial agent vancomycin was added to our panel; it is not included in the recommendations of the approved standard M24-A.61 Since no interpretive criteria for the breakpoints for vancomycin have been recommended for Gordonia species, those for bacteria that grow aerobically were used.62 Our susceptibility studies of 23 available type strains are shown in Table 9.2. All species tested were susceptible to amikacin, imipenem, linezolid, and minocycline; all species were susceptible to vancomycin except G. effusa, which was intermediate; all species were susceptible to clarithromycin except G. bronchialis; all species were susceptible to ciprofloxacin except G. otitidis and G. sputi; most species were susceptible to amoxicillin/clavulanate except G. sinesedis, G. soli, and G. terrae; and most species were susceptible to ceftriaxone except G. nambiensis, G. sihwensis, and G. soli. More than half (61%) of the type strains of Gordonia species were resistant to trimethoprim/ sulfamethoxazole. Until recently, there were few studies on the antimicrobial susceptibility data of clinical isolates of Gordonia. Comparisons of our study with these studies are difficult because either no type strains were included or the species identification was not given. In 2007, Blaschke et al.39 reported their susceptibility studies (n = 6) as well as data that were previously published for a total of 24 isolates. The authors deducted from their analysis that initial therapy for Gordonia could include a carbapenem or a fluoroquinolone in combination with an aminoglycoside. These authors also noted that 35% of the isolates were resistant to trimethoprim/ sulfamethoxazole (often used to treat Nocardia infection) and 11% of the isolates were resistant to vancomycin (often used for treatment of Rhodococcus infection).39 Our results

S

Color

O

−

2

−

−

−

−

−

+

−

+

+

−

−

+

−

+

−

−

−

−

−

+

+

−

+

+

−

Adonitol

l-Arabinose

Cellobiose

Dulcitol

i-Erythritol

d-Fructose

d-Galactose

d-Glucose

Glycerol

i-myo-Inositol

Lactose

Maltose

d-Mannitol

Mannose

Melibiose

Raffinose

l-Rhamnose

Salicin

d-Sorbitol

Sucrose

Trehalose

d-Xylose

Growth at 25°C

35°C

45°C

−

+

+

−

+

+

−

−

−

−

−

+

−

+

−

−

+

W+

−

+

−

−

−

−

−

Assimilation w/acid production

−

Acid-Fastb

Characteristic

1

−

+

+

−

+

+

−

+

+

−

−

+

+

+

−

+

−

+

−

+

−

−

−

−

−

Y

−

3

−

+

+

−

+

+

+

−

−

−

−

−

+

−

−

−

+

+

−

+

−

−

−

−

−

O

+

4

−

+

+

−

+

−

−

+

−

−

−

+

−

−

−

−

+

+

−

+

−

−

−

−

−

W

+

5

−

+

+

−

+

+

−

−

−

−

−

+

−

−

−

+

+

+

−

+

−

−

−

−

−

O

−

6

NT

NT

NT

NT

NT

NT

NT

NT

NT

NT

NT

NT

NT

NT

NT

NT

NT

NT

−

NT

NT

NT

NT

NT

NT

B

+

7a

NT

NT

NT

NT

NT

NT

NT

−

−

NT

NT

−

NT

−

NT

NT

−

NT

−

−

NT

NT

−

−

NT

NP

+

8a

−

+

+

−

+

+

−

−

−

−

−

+

−

−

−

−

+

+

−

+

−

−

−

−

−

B

−

9

−

+

+

−

+

+

−

−

−

−

−

+

+

−

−

−

+

+

−

+

−

−

−

−

−

B

−

10

−

+

+

−

+

+

−

−

−

−

−

+

−

−

−

+

+

+

−

+

−

−

−

−

−

Y

−

11

TABLE 9.1 Phenotypic Characteristics of the Type Strains of the Genus Gordonia

−

+

+

−

+

+

−

−

−

−

−

−

−

+

−

−

−

+

−

+

−

−

−

−

−

Y

+

12

NT

NT

NT

NT

NT

+

NT

NT

−

NT

NT

NT

NT

NT

NT

+

NT

NT

−

NT

NT

NT

NT

NT

NT

W

NT

13a

NT

NT

NT

+

+

+

NT

W

+

W

W

+

+

NT

+

W

NT

+

W

+

NT

NT

+

+

W

P/O

+

14a

NT

NT

NT

+

+

+

−

NT

−

−

NT

NT

−

−

−

−

NT

+

−

NT

−

NT

−

−

−

C

−

15a

−

+

+

−

+

+

+

−

−

−

−

+

+

−

−

−

+

+

−

+

−

−

−

−

−

O

−

16

−

+

+

+

+

+

+

−

−

−

−

+

+

−

−

−

+

+

−

+

−

−

−

−

−

S

+

17

+

+

+

−

+

+

−

−

−

−

−

+

−

−

−

−

+

+

−

−

−

−

−

−

−

O

−

18

−

+

+

+

+

+

+

−

+

−

−

+

+

−

−

+

+

+

+

+

+

−

−

+

+

Y

−

19

−

+

+

−

+

+

−

−

+

−

−

+

+

+

−

+

+

+

−

+

−

−

+

−

−

S

−

20

−

+

+

+

+

+

+

−

+

−

−

+

+

−

−

+

+

+

+

+

−

−

−

−

+

O

+

21

−

+

+

−

+

−

−

−

−

−

−

−

−

+

−

−

−

+

+

−

−

−

−

−

−

Y

−

22

+

+

+

−

+

+

−

−

−

−

−

+

−

−

−

−

+

+

−

+

−

−

−

−

−

Y

−

23

+

+

+

−

+

+

−

−

−

−

−

+

−

+

−

+

+

+

−

+

−

−

−

−

−

Y

+

24

−

+

+

−

+

+

+

−

−

−

−

+

+

+

−

−

+

+

−

+

−

−

−

−

−

O

−

25

−

+

+

−

+

+

+

−

−

−

−

+

+

+

−

−

+

+

−

+

−

−

−

−

−

O

+

26

−

+

+

+

+

+

−

+

+

+

+

+

+

+

+

−

+

+

+

+

−

+

+

+

−

S

+

28

continued

−

+

+

−

+

+

+

−

−

−

−

−

−

+

−

−

−

+

−

+

−

−

−

−

−

O

−

27

Gordonia 99

+

Nitrate reduction

+

−

+

−

−

−

+

+

−

−

+

−

3

−

−

−

−

−

−

−

−

−

−

4

+

+

−

−

−

−

−

−

+

−

5

−

−

−

+

+

+

−

−

−

−

6

NT

NT

NT

−

NT

NT

NT

NT

NT

NT

7a

+

NT

NT

NT

−

NT

−

−

−

NT

8a

−

−

−

+

+

+

−

−

−

−

9

+

−

+

+

−

+

−

−

+

−

10

+

−

−

−

−

−

−

−

−

−

11

+

−

+

+

+

+

−

−

−

−

12

−

NT

NT

−

−

+

NT

NT

+

−

13a

+

NT

NT

+

NT

W

−

−

−

+

14a

NT

NT

NT

+

−

+

−

−

−

−

15a

+

−

−

+

−

+

−

−

−

−

16

+

+

+

+

+

+

−

−

−

−

17

+

−

−

+

+

+

−

−

−

−

18

−

−

+

+

+

+

−

−

−

−

19

+

+

−

+

−

+

−

−

−

−

20

−

−

−

+

+

+

−

−

−

−

21

+

−

+

−

+

−

−

−

−

−

22

+

−

−

+

+

−

−

−

−

−

23

+

−

+

+

−

+

+

+

−

−

24

+

−

−

+

−

+

−

−

−

−

25

+

−

−

−

−

+

−

−

+

−

26

+

−

−

+

−

+

−

−

−

−

27

+

−

−

+

+

+

−

+

+

−

28

Notes: 1, G. aichiensis; 2, G. alkanivorans; 3, G. amarae; 4, G. amicalis; 5, G. araii; 6, G. bronchialis; 7, G. cholesterolivorans30; 8, G. defluvii25; 9, G. desulfuricans; 10, G. effusa; 11, G. hirsuta; 12, G. hydrophobica; 13, G. kroppenstedti31; 14, G. lacunae29; 15, G. malaquae28; 16, G. namibiensis; 17, G. otitidis; 18, G. paraffinivorans; 19, G. polyisoprenivorans; 20, G. rhizosphera; 21, G. rubripertincta; 22, G. shandongensis; 23, G. sihwensis; 24, G. sinesedis; 25, G. soli; 26, G. sputi; 27, G. terrae; 28, G. westfalica. All species showed a rod to cocci microscopic morphology, except for G. cholesterolivorans and G. defluvii; these two species additionally showed hyphae. Abbreviations and symbols:  −, negative; +, positive; NT, not tested; S, salmon; O, orange; Y, yellow; W, white; B, beige; NP, no pigment; P, pink; C, cream; W+, weak positive. a Assimilation/utilization data from previously published original manuscripts; not acid production. b Acid-fast organisms usually from 4 to 10 per oil immersion field.

−

Arylsulfatase production

−

+

−

+

Citrate

+

Growth in lysozyme

Acetamide

+

−

−

+

−

2

+

+

Urea

Utilization of

−

−

Hypoxanthine

+

Esculin

Tyrosine

−

Casein

Hydrolysis of

1

TABLE 9.1 (Continued) Phenotypic Characteristics of the Type Strains of the Genus Gordonia

100 Molecular Detection of Human Bacterial Pathogens

S S

S S

DSM 44369

ATCC 27808

DSM 44461

G. amarae

G. amicalis

S S

ATCC 14352

G. rubripertincta

S S

ATCC 25594

DSM 44215

G. terrae

G. westfalica

c

b

S

R

S

R

R

S

S

S

S

S

S

S

S

S

S

S

R

S

R

S

S

S

S

S

S

S

S

R

S

S

S

S

S

I

S

S

S

S

Ampicillin

S

S

S

R

S

R

S

S

S

S

S

S

R

S

S

S

S

S

S

S

S

S

S

Ceftriaxone

S

S

S

S

S

S

S

S

S

S

S

S

S

S

S

S

S

R

S

S

S

S

S

Clarithromycin

S

S

R

S

S

S

S

S

S

S

S

R

S

S

S

S

S

S

S

S

S

S

S

Ciprofloxacin

Antimicrobial Agent

S

S

S

S

S

S

S

S

S

S

S

S

S

S

S

S

S

S

S

S

S

S

S

Imipenem

Abbreviations: amox/clav, amoxicillin/clavulanic acid; TMP/SMX, trimethoprim/sulfamethoxazole; S, susceptible; I, intermediate; R, resistant. Based on interpretations of MICs, using CLSI breakpoints.61 Interpretive breakpoints for vancomycin are derived from M7-A7, CLSI.62

S

ATCC 29627

G. sputi

a

S S

DSM 44455

DSM 44995

G. sinesedis

G. sihwensis

G. soli

S S

JCM 13907

DSM 44576

G. shandongensis

G. polyisoprenivorans S

S

DSM 44604

G. paraffinivorans

DSM 44383

S

DSM 44809

G. otitidis

ATCC BAA-14

S

DSM 44568

G. rhizosphera

S

ATCC 700089

G. hydrophobica

G. namibiensis

S

S S

IFM 10200

ATCC 700255

G. effusa

S

DSM 44462

G. desulfuricans

G. hirsuta

S

S

S

S

IFM 10211

ATCC 25592

G. araii

G. bronchialis

S

S

S

S S

ATCC 33611

G. aichiensis

G. alkanivorans

Amox/ Clav

Amikacin

Species

Type Strain No.

TABLE 9.2 Antimicrobial Susceptibility Results of Type Strains of the Genus Gordoniaa,b

S

S

S

S

S

S

S

S

S

S

S

S

S

S

S

S

S

S

S

S

S

S

S

Linezolid

S

S

S

S

S

S

S

S

S

S

S

S

S

S

S

S

S

S

S

S

S

S

S

Minocycline

R

R

R

R

R

R

R

R

S

R

R

R

S

S

S

R

S

R

R

S

S

S

S

TMP/ SMX

S

S

S

S

S

S

S

S

S

S

S

S

S

S

S

I

S

S

S

S

S

S

S

Vancomycinc

Gordonia 101

102

were comparable to the studies of Aoyama et al.34 except that in their report all 31 strains of clinical Gordonia had MICs of >2/38 for trimethoprim/sulfamethoxazole that were incorrectly interpreted as susceptible. This incorrect interpretation has been corrected (Mikami, Y., personal communication, 2009). Although Aoyama et al.34 determined susceptibility studies for 31 molecularly identified strains (14 G. sputi, 13 G. bronchialis, two strains of G. aichiensis, and one strain each of G. otitidis and G. terrae); these authors did not use any type strains. In another series of nine clinical isolates (five isolates of G. terrae and four isolates of G. sputi) by Lai et al. in 2009,51 results were comparable to our study of 23 type strains except Lai et al. found all nine isolates were susceptible to trimethoprim/sulfamethoxazole.51 These discordant results may reflect different testing methods or difficulties in the interpretation of breakpoints for trimethoprim/ sulfamethoxazole. 9.1.5.2  M  olecular Techniques for Identification and Phylogenetic Analysis Need for Molecular Identification Techniques. In the past two decades, the need by clinical and reference laboratories for a rapid and accurate method for identifying Gordonia clinical isolates has increased due to the rising number of reports of human infections in part as well as an increasing awareness of these infections by clinicians. Identification of Gordonia, as well as for other clinically or environmentally relevant bacteria, using conventional biochemical and physio­logic methods is difficult due to the potentially long incubation times needed for identification, the availability of only a few discriminatory tests, and the high degree of expertise required to perform the tests.63 Gordonia clinical isolates have often been misidentified or regarded as nocardioform actinomycete contamination when cultures grew Gordonia. Accurate identification of patient specimens is important for understanding its epidemiology, identifying its clinical importance, defining its spectrum of disease, assessing its pathogenicity, and establishing risk factors for infection. Molecular methods for identification, especially DNA sequencing, have become increasingly an important adjunct to conventional phenotypic identification methods. A polyphasic approach provides accurate and reliable identification for existing and newly characterized species. DNA sequencing generally provides a more rapid and definitive identification than can be obtained by biochemical analysis alone. Molecular characterization of bacteria may also provide clinically relevant information on the phylogenetic relationship between pathogenic Gordonia species and other bacteria. Molecular identification has been aided by technological advances in DNA sequencing. These advances include the availability of automated sequencers, commercially available reagents, improved software for analysis, and more comprehensive and reliable gene databases. DNA RFLP Analysis. Restriction fragment length polymorphism (RFLP) analysis provided an early method for molecular identification of Gordonia clinical isolates. Briefly, a 439 bp DNA fragment of the 65 kDa heat shock protein

Molecular Detection of Human Bacterial Pathogens

gene (groE) was amplified by PCR. The PCR product was digested with a battery of restriction endonucleases and the restriction fragments were resolved by MetaPhor agarose gel electrophoresis. Following electrophoresis, gels were stained and identification was made by comparing the restriction fragment profiles obtained from the clinical isolate to the profiles of other type or reference strains.64 PCR–RFLP was able to distinguish between the profiles obtained from clinical isolates of actinomycetes, including G. bronchialis and G. sputi. However, interpretation was difficult due to multiple patterns obtained from isolates of G. bronchialis. RFLP analysis of the 16S rRNA gene-containing fragments or ribotype analysis has also been used for the identification of Gordonia clinical isolates. Ribotyping starts by digestion of purified genomic DNA with a restriction endonuclease. The DNA fragments are separated by agarose gel electrophoresis and the DNA fragments are transferred by the method of Southern to a solid support such as a nitrocellulose or nylon filter. Southern blots are hybridized with a labeled 16S rRNA probe. Identification is made following comparison of the RFLP hybridization profiles of 16S rRNA gene-containing fragments obtained from the unknown isolate to the hybridization profiles obtained from reference or type strains.65,66 Ribotyping was used by Buchman et al.65 as an adjunct method to identify an isolate obtained from a case patient with central venous catheter sepsis since traditional biochemical identification was ambiguous between G.  rubripertincta and G. terrae. The profile of 16S rRNA gene-containing fragments was similar to the rRNA gene fragment profile obtained from the G. terrae type strain and the profile clearly differed from the 20 other type strains. However, identification of the isolate from a second patient using ribotyping was not reliable due to heterogeneity of the profiles for G. sputi and G. bronchialis isolates. Ribotype analysis has been used as an adjunct method for identification of G. terrae in a brain abscess52 and in a mycetoma patient in Sierra Leone.57 Ribotyping has been successful as an adjunct method for identification and typing of a cluster of patient isolates identified as G. bronchialis obtained from sternal wound infections after coronary bypass surgery.35 In addition, ribotyping confirmed the results obtained by DNA– DNA hybridization, and was instrumental in the reevaluation of the taxonomic position of Rhodococcus chubuensis to G. sputi.67 Gordonia environmental isolates have been identified by use of PCR-denaturing gradient gel electrophoresis (DGGE).68–70 Briefly, Gordonia-specific primers, G268F and G1096R, were used to amplify 16S rRNA genes by PCR. rRNA-containing bands were either excised from agarose gels for direct DNA sequencing or the excised sample was further resolved by DGGE analysis. Following DGGE, bands were excised and subjected to DNA sequence analysis. Identification is made by comparing 16S rRNA gene sequence obtained from the unknown organism to DNA sequences within an appropriate sequence database with the high degree of sequence similarity. PCR–DGGE has detected and identified Gordonia in oil-contaminated soils  and  environmental

Gordonia

samples  but  has  never  been applied to the identification of clinical samples. DGGE analysis is more sensitive than conventional methods for recovery of rRNA gene sequences; however, the extra steps required for DGGE increase the risk of cross contamination and may be too technically demanding for most clinical laboratories. PCR–RFLP and ribotyping have been previously useful for identification, but have many weaknesses. First, there are problems associated with ensuring complete digestion of genomic DNA and accurate size determination of restriction fragments. For example, each sample should be run with internal standards to correct for lane-to-lane or between run variability that would make intraand interlaboratory comparisons especially difficult. A  second obstacle is the lack of a standard protocol and criteria for interpretation. Interpretation of profiles is critical since variant profiles may be obtained by a new species or by intraspecies heterogeneity. Third, due to the increasing number of restriction endonucleases that would be required to resolve all validated species and recognize newly recovered species, the method rapidly becomes more complex. As may be expected, failure of the method to reliably identify Gordonia clinical isolates has previously been documented.48,53 DNA Sequencing. Of all the gene targets found useful for identification and or phylogenetic analysis of bacteria, 16S rRNA gene sequences are used most widely. Their widespread utilization is due to several factors. First, the 16S rRNA gene is universally present in all bacteria and the biological function of the gene is well known. Second, 16S rRNA genes evolve at a slower rate relative to that observed for house keeping genes. The slower rate is due to the constraints of the three-dimensional structure required for a functional 16S rRNA molecule. Therefore, the gene includes both conserved and variable sequence motifs. Conserved sequence motifs may provide consensus family- or genusspecific sequences whereas variable regions may provide for species-specific identification. The 16S rRNA gene has been found to be appropriate for phylogenetic analysis due to these slowly evolving conserved sequences. Lastly, the gene is relatively easy to sequence and its length—approximately 1.4 kb—has been determined to be large enough for phylogenetic analysis. Both clinical and environmental isolates of Gordonia spp. have been identified using 16S rRNA gene sequences. Isolates of G. aichiensis and G. amarae were among the first pathogenic strains to be identified and classified using analysis of 16S rRNA gene sequences.8,71 To expedite the rapid detection and identification of Gordonia species in complex environmental samples, Patel et al.72 developed primer pair, G268F and G1096R, to amplify by PCR an 828 bp 16S rRNA gene fragment.68 The primer pair was able to amplify the appropriate sized fragments from 19 Gordonia type strains and Gordonia-specific signature nucleotides were observed. The primer pair may be Gordonia-specific since it failed to amplify the 16S gene for other species of actinomycetes. Analysis of 16S rRNA gene sequences for 47 strains of Gordonia, including pathogenic strains of G. bronchialis, G. terrae, G. polyisoprenivorans, and G. sputi, formed a distinct clade that was separate from

103

phylogenetically related actinomycete species. However, not all species of Gordonia were distinguishable using 16S rRNA gene sequences. Reliable identification was not always possible since the fragment length was short, at 828 bp. Longer 16S rRNA gene sequences have been shown to improve the accuracy and reliability of molecular identification for other species of bacteria.73 Identification of clinical isolates by 16S rRNA gene analysis is an increasingly important adjunct method to tradition biochemical diagnostics and polyphasic characterization. However, there are several difficulties that may impede accurate identification. For instance, public and private sequence databases have been known to harbor sequences containing errors, ambiguous bases, or misidentified strains due to lack of quality control or a protocol to remove errors. The collection of reference and type species may be either incomplete or requires updating. A second problem is the lack of a universal standard for interpretation, especially for identification of species with nearly identical 16S sequences. Simply, how similar should sequences be for reliable and accurate identification? Many species have recently evolved and have nearly indistinguishable sequences. These recently evolved species are difficult to identify due to lack of sequence divergence in a short time period or gene sequences made homogeneous due to recombination. For example, the sequence of a 450 bp 16S rRNA gene fragment from a patient with bacteremia and endocarditis was 100% identical to the 16S rRNA gene sequence of G. sputi but the relatedness of the isolate to G. sputi by DNA–DNA hybridization analysis was only 55%, far below the threshold of 70% recommended for the definition of same species.56 Likewise, the nearly full-length 16S rRNA gene sequence obtained from a patient with bacteremia differed by the 16S rRNA gene sequence of G. sputi and G. aichiensis by 1 and 3 bp, respectively, whereas the isolate was determined to be 60% or 21% related by DNA–DNA hybridization analysis to G. sputi or G. aichiensis, respectively.48 These investigations demonstrate the practical difficulties of reliable identification of Gordonia spp. based solely on analysis of 16S rRNA gene sequences. Accurate identification may be mired by the presence of multiple copies of 16S rRNA genes resulting in ambiguous bases or sequence heterogeneity since the majority rule consensus sequence ignores real allelic differences. Other issues involve the inability to identify samples obtained from mixed cultures, the high cost of supplies, technical expertise, and equipment, all of which may be a significant factor delaying procurement of routine DNA sequencing by many clinical laboratories. To circumvent the difficulties associated with analysis of 16S gene sequences, an alternative gene target, DNA gyrase subunit B (gyrB), was sequenced and directly compared to analysis using 16S rRNA gene sequences for identification and phylogenetic analysis of Gordonia isolates.74 Briefly, genomic DNA was purified and a ~1.2 kb gyrB gene fragment was amplified by PCR using primers GYRB1 and GYRB2 for 25 different strains representing 17 different type strains of Gordonia. GyrB amino acid and gyrB nucleotide sequences were aligned by Clustal X software and analyzed

104

using MEGA 2.1 software. The range of sequence similarity for 12 Gordonia type strains gyrB nucleotide sequences was 79.3%–97.2%, corresponding to 270–41 bp differences. In comparison, a sequence difference of 0.3%–3.8% was observed by 16S rRNA gene sequencing for the same 12 type strains. The branching orders of the 16S rRNA and gyrB gene trees were similar, but gyrB gene sequences showed a higher degree of separation between species. Enhanced discrimination may be due to the higher degree of nucleotide substitution rate observed for the gyrB gene than observed for 16S rRNA gene sequences. Interestingly, the correlation coefficients of GyrB amino acid sequences with DNA–DNA homology and 16S rRNA gene sequences with DNA–DNA homology were both identical (R2 = 0.865). Nucleotide substitutions were scattered over the entire length of the gyrB gene. In contrast, nucleotide substitutions in the 16S rRNA gene were concentrated to a few variable regions, thereby allowing gyrB to provide better discrimination between species than obtained by 16S rRNA gene sequences. In summary, Shen et al.74 reported gyrB gene sequences were a good alternative and appropriate marker for identification of Gordonia species. In a similar investigation, Kang et al.75 compared the phylogenetic relationships obtained between 23 Gordonia type strains using three different gene sequences: gyrB, secA1, and 16S rRNA. Kang et al. amplified an approximate 1230 bp gyrB gene fragment using PCR primer pairs, F1 and R1296, and fragments were then sequenced. Similarity of the GyrB amino acid sequence and gyrB gene sequence ranged from 75.7% to 99.8%, and 77.5% to 97.6%, respectively. The gyrB gene tree branching order obtained by Kang et al.75 was consistent with the investigation by Shen et al.68 A 496 bp gene fragment, representing approximately 160 amino acids of the secA1 gene, was amplified by PCR and sequenced. Interspecies sequence similarity for the secA1 gene sequence and the SecA1 amino acid sequence was 81.9%–98.0%, and 83.0%–100%, respectively. Additionally, a secA1 signature sequence for Gordonia species, IKKK, located between amino acid residues 120–130, was identified. In comparison, the interspecies sequence similarity for 16S rRNA gene sequences ranged from 94% to 99.8%; this is higher than obtained for gyrB and secA1 gene sequences. Most of the nucleotide variability in the 16S rRNA gene resides between nucleotide positions (based on Escherichia coli strain K-12) 136–229 and 996–1028.3 The branching order for the three gene trees was highly similar but not identical, suggesting a variable rate of evolutionary change for each gene. Both gyrB and secA1 gene sequences provided a higher degree of discrimination than that obtained by 16S rRNA gene sequences, whereas the highest degree of discrimination and identification was observed with gyrB gene sequences. Phylogenetic analysis using the combination of these three gene sequences allowed for the unambiguous identification and resolution of all 23 Gordonia type strains and resulted in a higher degree of resolution obtainable than by using a sequence from a single gene. Kang et al.75 concluded that both gyrB and secA1 gene sequences

Molecular Detection of Human Bacterial Pathogens

are appropriate molecular markers for identification and phylogenetic analysis of Gordonia species isolates. At present, only a few clinical or reference laboratories utilize gyrB and secA1 gene sequences for identification of Gordonia clinical isolates. One difficulty is that not all species of Gordonia are capable of being amplified using a single standard amplification regimen with primers F1 and R1296.75 Other difficulties include establishment of a similarity cutoff criteria for reliable and accurate identification since intraspecies sequence variability has been observed. For example, the 16S rRNA gene similarity ranged from 99.6% to 99.9% for six isolates of G. terrae. However, analysis of gyrB gene sequences demonstrates two subclades. One isolate has 98.0% sequence similarity to the type strain whereas the sequence similarity for the remaining four isolates was 87.0%, suggesting that G. terrae may be heterogeneous, composed of a physiological similar but genetically different set of isolates. DNA–DNA hybridization or analysis of other molecular markers may be required to determine whether G. terrae is a heterogeneous species. Identification of Clinical Isolates Using DNA Sequencing. There are several reports of Gordonia clinical isolates misidentified by routine biochemical tests such as API Coryne or by other phenotypic tests that rely primarily on the analysis of 16S rRNA gene sequences for accurate identification.33,36,38,39,43,54,76 For instance, an isolate obtained from a patient with a sequestrated lung34 and from a patient with a recurrent breast abscess38 were incorrectly identified as a Rhodococcus species by the API Coryne, but were identified correctly as G. bronchialis by analysis of 16S rRNA gene sequences. Similarly, a blood culture isolate from a patient with endocarditis was initially identified as a Corynebacterium. This isolate was later identified as G. polyisoprenivorans since 1444 of 1445 bp of the 16S rRNA gene sequences were identical to the G. polyisoprenivorans type strain.43 In a large retrospective study in Japan, 14 of 34 clinical isolates of Gordonia identified by biochemical profiles were then confirmed by analysis of 16S rRNA gene sequences as G. sputi34; however, these isolates may be heterogeneous since 16S rRNA gene analysis detected up to four genetic groups. G. sputi, identified in sputum samples using morphologic and biochemical tests (before the availability of routine 16S gene sequencing), were considered a source of pulmonary infections.45,49,63 However, it may be interesting to reevaluate these studies with 16S rRNA gene sequences to provide a more robust and accurate identification. A clinical isolate of G. araii, associated with an orthopedic device in an immunocompetent patient, used both 16S rRNA and gyrB sequences for identification, since biochemical analysis was unreliable for accurate identification.33 Analysis of 16S rRNA gene sequences may also be used for the initial identification and characterization of novel species of Gordonia. For instance, two recently described pathogenic species of Gordonia, G. araii, and G. effusa, isolated from sputum of patients with bacterial pneumonia or kidney dysfunction, respectively, were identified based on differences in 16S rRNA gene similarity to other known Gordonia species.24

105

Gordonia

Similarly, analysis of 16S rRNA gene was used for definitive identification of a strain obtained from an ear discharge and was later identified as a new species, G. otitidis.23 Despite the known limitations of solely using 16S rRNA gene analysis for species identification, routine use of these gene sequences presently provides a useful and practical tool to identify rare or unknown clinical isolates. Microarrays and Whole Genome Sequencing. Microarrays or microchips generally consist of a complex mixture of short, specific oligonucleotide sequences immobilized on a solid support. The immobilized oligonucleotides are hybridized with a mixture of labeled nucleic acid probes allowing simultaneous detection and quantitative measurement of a complex mixture of specific gene sequences. DNA microarrays have the potential to be used for identification of pathogenic Gordonia or other bacteria. Microarrays may also be fabricated to determine antimicrobial resistance or detect patterns of transcription during infection. As a proof of concept for rapid species identification using microarray technology, a microarray was fabricated with 45 random, unsequenced, genomic DNA sequences of unknown function. All were 200–1500 bp in length and were obtained from G. amarae genomic DNA.77 G. amarae microarrays were hybridized with labeled genomic DNA, purified from thirteen different species of bacteria or DNA purified from G. amarae itself. The microarray was observed to accurately identify G. amarae since positive signals were obtained only from G. amarae DNA and showed no cross hybridization by other species of bacteria. Positive signals were also observed using G. amarae DNA mixed with heterologous DNA probe. Sequencing of the entire Gordonia species genomes or particular gene sequences may provide species-specific sequences useful for the fabrication of Gordonia-specific microarrays.

9.2  METHODS 9.2.1  Sample Preparation Presently, molecular identification of Gordonia isolates requires culturing of clinical samples on appropriate growth medium since DNA used for molecular analysis is obtained from pure cultures. Clinical samples are commonly cultured aerobically on rich medium including trypticase soy agar, brain heart infusion agar, and chocolate or blood agars. Species of Gordonia have been isolated from a vast array of clinical samples including blood,39,42,43,48,55,56 sputum,34,45,63 biopsied tissue,33,38,46,57,58 CSF,39,53,54 and a lymph node.47 Nucleic acids for molecular analysis are purified from colonies grown on the appropriate solid agar or obtained from cells grown in broth. Cells may be lysed and DNA purified using a combination of enzymatic digestion and deproteinization by repeated phenol-chloroform-isoamyl alcohol extractions; lysis using sodium dodecyl sulfate (SDS) in the presence of proteinase K35; or lysis with guanidine thiocyanate.23,24 DNA has been obtained by cell rupture using repeated cycles of freezing in liquid nitrogen and boiling53 or with a beadbeater using zirconium beads.64 Recently, high

purity DNA has been obtained using commercially available DNA isolation kits such as the soil DNA Extraction Kit,78 the UltraClean Soil DNA Isolation Kit,69,78 the UltraClean Microbial Genomic DNA Isolation Kit,74 the ISOPLANT I Kit,75 or the DNAeasy Tissue Kit.36

9.2.2  Detection Procedures (i) Phylogenetic analysis of 16S rRNA gene seque­ nces  Phylogenetic relationships among Gordonia species isolates can be determined following the amplification, alignment, and analysis of near full-length 16S rRNA gene sequences. Procedure 1. Amplification of near full-length 16S rRNA gene is performed in a 50 μL reaction mixture consisting of approximately 10 ng of purified genomic DNA, 200 μM of each dNTP, 5% (vol/vol) dimethyl sulfoxide, 2.5 U of Expand High-Fidelity DNA Polymerase (Roche Diagnostics Corp), and 100 nM each of primers fD1 (5′-CCGAATTCGTCG ACAACAGAGTTTGATCCTGGCTCAG-3′) and rP2 (5′-CCCGGGATCCAAGCTTACGGCTACC TTGTTACGACTT-3′) in the buffer supplied by the manufacturer. 2. Amplification conditions consist of an initial denaturation step at 94°C for 5 min followed by 35 cycles of amplification at 94°C for 15 s, 50°C for 15 s, and 72°C for 1.5 min, with a final extension at 72°C for 5 min. 3. PCR products are confirmed by visualization on 1.2% agarose gels. 4. Primers and nucleotides are removed with ExoSap-It (USB Corporation) using the reagents and protocol supplied by the manufacturer. 5. Purified amplicons are then sequenced in the forward and reverse directions by cycle sequencing with the ABI Prism BigDye terminator cycle sequencing ready reaction kit (Perkin Elmer/Applied Biosystems). Sequencing reactions are then loaded and resolved by capillary electrophoresis using an ABI 3730 DNA analyzer (Applied Biosystems). 6. Chromatograms are edited, assembled, and trimmed using Seqmerge software (Wisconsin Package version 10.3; Accelrys, Inc., San Diego, CA). 7. High-quality consensus sequences can be submitted to GenBank for comparative analysis using BLASTn software available at the BLAST Web site http://www.ncbi.nlm.nih.gov/BLAST. Since there is no universally recognized criterion for identification to the genus or species levels using 16S rRNA gene sequences, isolates were identified using the identity of the type strain with the highest degree of sequence similarity. 8. Phylogenetic analysis of 16S rRNA gene sequences is performed following alignment of nucleotide

106



Molecular Detection of Human Bacterial Pathogens

sequences using CLUSTAL W software. The aligned sequences are then used to construct a neighbor-joining tree using MEGA software. 9. Tree topology is evaluated by bootstrap analysis based on 1000 resamplings using MEGA software. Additional 16S rRNA primers for phylogenetic analysis of Godonia spp. are listed in Table 9.3.

(ii) Phylogenetic analysis using gyrB gene seque­ nces  Phylogenetic relationships among Gordonia species isolates can be determined following the alignment and analysis of gyrB gene sequences. Procedure 1. Amplification of the DNA gyrase β-subunit (gyrB) are performed using 20 ρmol of primers GYRB1 (5′-ATGACNCARYTNCAYGCNGGN-3′) and GYRB2 (5′-SAYGATCTTGTKRTASCGMAAYT T-3′) in a volume of 20 μL reaction mixture using the protocol and amplification conditions described by Shen et al.74 Alternatively, amplification was performed with 20 ρmol each of primers GYRB1 and GYRB2, 0.2 mM of each of the four dNTPs, 5% (vol/vol) dimethyl sulfoxide, and 0.2 U of iProof High-Fidelity DNA Polymerase in iProof GC buffer (BioRad).







2. Amplification conditions consisted of an initial denaturation of template DNA at 98°C for 1 min, followed by 30 cycles, consisting of 98°C for 15 s, 59°C for 30 s, and 72°C for 1 min, followed by a final extension at 72°C for 5 min. 3. PCR products are confirmed by visualization on 1.2% agarose gels. 4. Primers and nucleotides are removed with ExoSapIt (USB Corporation) using the reagents and protocol supplied by the manufacturer. 5. Purified amplicons are then sequenced in the forward and reverse directions by cycle sequencing with the ABI Prism BigDye terminator cycle sequencing ready  reaction kit (Perkin Elmer/ Applied Biosystems) using sequencing primers GYRB1, GYRBF1, GYRBF2, GYRB2, and GYRBR1, described previously by Shen74 and a new reverse sequencing primer,  R2 (5′-GAGATG GCGTTCCTSAACAAG-3′). Sequencing reactions are then loaded and resolved by capillary electrophoresis using an ABI 3730 DNA analyzer (Applied Biosystems). 6. Chromatograms are edited, assembled, and trimmed using Sequencher version 4.8 software (Gene Codes Corp, Ann Arbor, MI). 7. High-quality consensus sequences can be submitted to GenBank for comparative analysis using BLASTn

TABLE 9.3 16S rRNA Gene Primers for Amplification and Sequencing Gordonia spp. Primer fD1 rP2 fD1-5p F27 F785 R357 R802 F357 R530 R519 F530 BSF917 BSR926 BSF1099 BSR1114 BSF1391 BSR1407 rP2-5p

Sequence (5′–3′) CCGAATTCGTCG ACAACAGAGTTTGATCCTGGCTCAG CCCGGGATCCAA GCTTACGGCTACCTTGTTACGACTT CCGAATTCGTCGACAACAG AGAGTTTGATCMTGGCTCAG GGATTAGATACCCTGGTA CTGCTGCCTCCCGTA TACCAGGGTATCTAATCC TACGGGAGGCAGCAG GTATTACCGCGGCTGCTG GWATTACCGCGGCKGCTG CAGCAGCCGCGGTAATAC GAATTGACGGGGRCCC CCGTCAATTYYTTTRAGTTT GYAACGAGCGCAACCC GGTTGCGCTCGTTRC TGTACACACCGCCCGTC GACGGGCGGTGTGTRC CCCGGGATCCAAGCTTAC

Application

Orientationa

Referenceb

PCR PCR Sequencing Sequencing Sequencing Sequencing Sequencing Sequencing Sequencing Sequencing Sequencing Sequencing Sequencing Sequencing Sequencing Sequencing Sequencing Sequencing

F R F F F R R F R R F F R F R F R R

Weisburg et al., 1991 Weisburg et al., 1991 Morey et al., 2006 Heuer et al., 1997 Weisburg et al., 1991 Weisburg et al., 1991 Weisburg et al., 1991 Weisburg et al., 1991 Weisburg et al., 1991 Weisburg et al., 1991 Weisburg et al., 1991 Wuyts et al., 2002 Wuyts et al., 2002 Wuyts et al., 2002 Wuyts et al., 2002 Wuyts et al., 2002 Wuyts et al., 2002 Wuyts et al., 2002

  F, forward; R, reverse.   Additional sources: Weisburg, W.G. et al., 16S ribosomal DNA amplification for phylogenetic study. J. Bacteriol., 173, 697, 1991. Morey, R.E. et al., Species-specific identification of Leptospiraceae of 16S rRNA gene sequencing. J. Clin. Microbiol., 44, 3510, 2006. Heuer, H. et al., Analysis of actinomycete communities by specific amplification of genes encoding 16S rRNA and gel-­electrophoretic separation in denaturing   gradients. Appl. Environ. Microbiol., 63, 3233, 1997. Wuyts, J. et al., The European database on small subunit ribosomal RNA. Nucleic Acids Res., 30, 183, 2002. a

b

Gordonia





software available at the BLAST Web site http:// www.ncbi.nlm.nih.gov/BLAST. Since there is no universally recognized criterion for identification to the genus or species levels using gyrB gene sequences, isolates are identified using the identity of the type strain with the highest degree of sequence similarity. 8. Phylogenetic analysis of gyrB gene sequences is performed following alignment of nucleotide sequences using CLUSTAL W software and then used to construct a neighbor-joining tree using MEGA software. 9. Tree topology is evaluated by bootstrap analysis based on 1000 resamplings using MEGA software.

9.3  C  ONCLUSIONS AND FUTURE PERSPECTIVES A method for identification of Gordonia in tissues would help to eliminate unnecessary culturing and shorten the length of time for identification. One potential method for identification in tissues is by fluorescence in situ hybridization (FISH). Briefly, species- and or genus-specific fluorescently labeled oligonucleotide probes are designed to specifically hybridize to 16S rRNA in cells due to their relative abundance. Cells are hybridized following a permeabilizing step using enzymatic or acid treatment of fixed cells. The specific fluorescence of target cells, visualized with a fluorescent microscope, readily distinguishes target cells from nontarget cells and debris. At present, FISH has been used to identify only G. amarae in environmental samples, including activated sludge at wastewater treatment plants.79,80 FISH may be a useful method for rapid identification, estimating cell viability, or direct detection in paraffin-embedded tissue samples. De Los Reyes et al.79 magnified the fluorescent signal for detection of Gordonia in environmental samples by simultaneous fluorescent antibody staining and FISH. Presently, FISH has not been used for identification of clinical specimens or tissues. While FISH may provide rapid identification, laboratories would require a complete array of species-specific probes and would require additional probes for new species. As discussed above, microarrays have the potential for rapid and specific identification and detection of Gordonia species isolates in either environmental or clinical samples. Species-specific and genus-specific oligonucleotide sequences may be acquired from whole genome sequencing projects or genome sequencing surveys. Microarrays can also be fabricated for investigations of gene expression during infection and disease prognosis, host interactions, identification of antibiotic resistance markers and resistant alleles, response to therapy, and obtaining transcription profiles during infection. Microarrays specific for G. polyisoprenivorans may be useful for determining the molecular mechanism for biofilm formation and propagation in catheter materials. There is an urgent need for a rapid, reliable, and accurate gene target for the molecular identification of pathogenic bacteria, including Gordonia species. Several potential gene targets are being investigated.81,82 Criteria for selecting a new molecular

107

target are that the target gene must be easy to amplify using a standard protocol, the target gene must be present in the bacteria of interest, the observed sequence variation must be high enough for species identification but not too conserved that species may artificially cluster together, and the target must be easy to interpret for unambiguous identification. Alternatively, a strategy using multilocus sequencing may provide better identification and robust phylogenetic signals than obtainable by analysis of a single target gene. The phylogenetic reconstruction and identification scheme using the combination of 16S rRNA, secA1, and gyrB gene sequences by Kang et al.75 demonstrates the promise of this approach. Multilocus sequencing analysis is more labor intensive and costly than analysis of a single gene sequence. The method does, however, have the advantage of increased reliability and accuracy for species identification and in determining the phylogenetic relatedness of known clinical isolates and in identifying new species. However, at present, the optimal gene targets or the number of genes is not known. Pyrosequencing may provide an alternative method to conventional gene sequencing and has been shown to be useful for identification of species phylogenetically related to Gordonia including Mycobacterium.83 Pyrosequencing is based on real-time detection of DNA synthesis using a mixture of four enzymes. During DNA synthesis, when a deoxynucleotide has been incorporated into a DNA strand by Klenow Polymerase, pyrophosphate is liberated. ATP sulfurylase converts the pyrophosphate to ATP, which is then used in a light-producing reaction by luciferase.84 Pyrosequencing is faster and simpler than conventional DNA sequencing since the method does not rely on gels or expensive dyes for resolution of bases. The short read length of approximately 100 bp does pose a serious disadvantage, but may be compensated by sequencing of hypervariable regions of a gene. Purification of DNA is considered to be the most laborintensive and tedious step in molecular identification, and is therefore the most likely step to be improved by automation. A simple and efficient method of purification of genomic DNA from clinical specimens or cultures would greatly facilitate down stream methods for molecular diagnostics and DNA–DNA hybridization analysis. Likewise, rapid advances in DNA sequencing technology, automated systems, and lower costs may allow in-house molecular diagnostics for most clinical laboratories, including the ability to assemble and annotate entire genomes. Sequencing of entire Gordonia genomes may provide further information on unique gene targets for identification or phylogenetic analysis, for potential mechanisms of antibiotic resistance and for the discovery of unique metabolic pathways. The frequent misidentification of Gordonia species suggests the potential underestimation and thus under reporting of these emerging pathogens. Accurate and rapid identification by molecular methods is important for a reliable description of pathogenesis, epidemiologic investigations, application of appropriate antimicrobial therapy, and patient management. Together, these results strongly suggest using a polyphasic approach of combining morphological, biochemical, and molecular analysis for strain identification.

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Molecular Detection of Human Bacterial Pathogens 19. Kim, K.K. et al., Gordonia sihwensis sp. nov., a novel nitratereducing bacterium isolated from a wastewater-treatment bioreactor, Int. J. Syst. Evol. Microbiol., 53, 1427, 2003. 20. Maldonado, L.A. et al., Gordonia sinesedis sp. nov., a novel soil isolate, Antonie van Leeuwenhoek, 83, 75, 2003. 21. Linos, A. et al., Gordonia westfalica sp. nov., a novel rubberdegrading actinomycete, Int. J. Syst. Evol. Microbiol., 52, 1133, 2002. 22. Xue, Y. et al., Gordonia paraffinivorans sp. nov., a hydrocarbon-degrading actinomycete isolated from an oil-producing well, Int. J. Syst. Evol. Microbiol., 53, 1643, 2003. 23. Lida, S. et al., Gordonia otitidis sp. nov., isolated from a patient with external otitis, Int. J. Syst. Evol. Microbiol., 55, 1871, 2005. 24. Kageyama, A. et al., Gordonia araii sp. nov. and Gordonia effusa sp. nov., isolated from patients in Japan, Int. J. Syst. Evol. Microbiol., 56, 1817, 2006. 25. Soddell, J.A. et al., Gordonia defluvii sp. nov., an actinomycete isolated from activated sludge foam, Int. J. Syst. Evol. Microbiol., 56, 2265, 2006. 26. Shen, F.T. et al., Gordonia soli sp. nov., a novel actinomycete isolated from soil, Int. J. Syst. Evol. Microbiol., 56, 2597, 2006. 27. Luo, H. et al., Gordonia shandongensis sp. nov., isolated from soil in China, Int. J. Syst. Evol. Microbiol., 57, 605, 2007. 28. Yassin, A.F. et al., Gordonia malaquae sp. nov., isolated from sludge of a wastewater treatment plant, Int. J. Syst. Evol. Microbiol., 57, 1065, 2007. 29. Le Roes, M., Goodwin, C.M., and Meyers, P.R., Gordonia lacunae sp. nov., isolated from an estuary, Syst. Appl. Microbiol., 31, 17, 2008. 30. Drzyzga, O. et al., Gordonia cholesterolivorans sp. nov., a cholesterol-degrading actinomycete isolated from sewage sludge, Int. J. Syst. Evol. Microbiol., 59, 1011, 2009. 31. Kim, K.K. et al., Gordonia kroppenstedtii sp. nov., a phenoldegrading actinomycete isolated from a polluted stream, Int. J. Syst. Evol. Microbiol., 59, 1992, 2009. 32. Lechevalier, M.P., and Lechevalier, H.A., Chemical composition as criterion in the classification of aerobic actinomycetes, Int. J. Syst. Bacteriol., 20, 435, 1970. 33. Jannat-Khah, D.P. et al., Gordonia araii infection associated with an orthopedic device and review of the literature on medical device-associated Gordonia infections, J. Clin. Microbiol., 47, 499, 2009. 34. Aoyama, K. et al., Characterization of clinical isolates of Gordonia species in Japanese clinical samples during 1998– 2008, Mycopathologia, 168, 175, 2009. 35. Richet, H.M. et al., A cluster of Rhodococcus (Gordona) bronchialis sternal-wound infections after coronary-artery bypass surgery, N. Engl. J. Med., 324, 104, 1991. 36. Sng, L.H. et al., Bacteremia caused by Gordonia bronchialis in a patient with sequestrated lung, J. Clin. Microbiol., 42, 2870, 2004. 37. Brust, J.C. et al., Five cases of bacteraemia due to Gordonia species, J. Med. Microbiol., 58, 1376, 2009. 38. Werno, A.M. et al., Recurrent breast abscess caused by Gordonia bronchialis in an immunocompetent patient, J. Clin. Microbiol., 43, 3009, 2005. 39. Blaschke, A.J. et al., Gordonia species: emerging pathogens in pediatric patients that are identified by 16S ribosomal RNA gene sequencing, Clin. Infect. Dis., 45, 483, 2007. 40. Linos, A., and Steinbuchel, A., Investigations of the microbial breakdown of natural and synthetic rubbers, DECHEMA Monogr., 133, 279, 1996.

Gordonia 41. Linos, A., and Steinbuchel, A., Microbial degradation of natural and synthetic rubbers by novel bacteria belonging to the genus Gordonia, Kautsch. Gummi. Kurstst., 51, 496, 1998. 42. Kempf, V.A. et al., Gordonia polyisoprenivorans septicemia in a bone marrow transplant patient, Eur. J. Clin. Microbiol. Infect. Dis., 23, 226, 2004. 43. Verma, P. et al., Native valve endocarditis due to Gordonia polyisoprenivorans: Case report and review of literature of bloodstream infections caused by Gordonia species, J. Clin. Microbiol., 44, 1905, 2006. 44. Gupta, M., et al., A rubber-degrading organism growing from a human body, Int. J. Infect. Dis., 12, e332, 2009. 45. Hart, D.H.L. et al., Lung infection caused by Rhodococcus, Aust. NZ J. Med., 18, 790, 1988. 46. Edwards, J.F., and Simpson, R.B., Nocardioform actinomycete (Rhodococcus rubropertinctus)-induced abortion in a mare, Vet. Pathol., 25, 529, 1988. 47. Tsukamura, M. et al., Mesenteric lymphadenitis of swine caused by Rhodococcus sputi, J. Clin. Microbiol., 26, 155, 1988. 48. Riegel, P. et al., Bacteremia due to Gordona sputi in an immunocompromised patient, J. Clin. Microbiol., 34, 2045, 1996. 49. Kuwabara, M. et al., Mediastinitis due to Gordona sputi after CABG, J. Cardiovasc. Surg., 40, 675, 1999. 50. Lesens, O. et al., Bacteremia and endocarditis caused by a Gordonia species in a patient with a central venous catheter, Emerg. Infect. Dis., 6, 382, 2000. 51. Lai, C.C. et al., Infections caused by Gordonia species at a medical center in Taiwan, 1997 to 2008, Clin. Microbiol. Infect., “Accepted Article” doi: 10.1111/j.1469-0691.200903333085.x 52. Drancourt, M. et al., Brain abscess in an immunocompromised child due to Gordona terrae: Case report and review of Gordona terrae infections, Clin. Infect. Dis., 19, 258, 1994. 53. Drancourt, M. et al., Gordona terrae central nervous system infection in an immunocompetent patient, J. Clin. Microbiol., 35, 379, 1997. 54. Gil-Sande, E. et al., Etiological misidentification by routine biochemical tests of bacteremia caused by Gordona terrae infection in the course of an episode of acute cholecystitis, J. Clin. Microbiol., 44, 2645, 2006. 55. Grisold, A.J. et al., Isolation of Gordonia terrae from a patient with catheter-related bacteraemia, J. Med. Microbiol., 56, 1687, 2007. 56. Pham, A.S. et al., Catheter-related bacteremia caused by the nocardioform actinomycete Gordonia terrae, Clin. Infect. Dis., 36, 524, 2003. 57. Bakker, X.R., Spauwen, P.H.M., and Dolmans, W.M.V. Mycetoma of the hand caused by Gordona terrae: A case report, J. Hand Surg., 29B, 188, 2009. 58. Zardawi, I.M. et al., Gordonia terrae-induced suppurative granulomatous following nipple piercing, Pathology, 36, 275, 2009. 59. Berd, D., Laboratory identification of clinically important aerobic actinomycetes, Appl. Microbiol., 25, 665, 1973. 60. Conville, P.S., and Witebsky, F.G., Nocardia, Rhodococcus, Gordonia, Actinomadura, Streptomyces, and other aerobic actinomycetes. In Murray, P. R., Baron, E. J., Jorgensen, J. H., Landry, M.I., and Pfaller, M.A. (Editors), Manual of Clinical Microbiology, 9th Ed., p. 515, American Society for Microbiology, Washington, D.C., 2007. 61. Clinical and Laboratory Standards Institute. Susceptibility testing of mycobacteria, nocardiae, and other aerobic actinomycetes. Approved standard M24-A. National Committee for Clinical Laboratory Standards, Wayne, PA, 2003.

109 62. Clinical and Laboratory Standards Institute. Methods for dilution antimicrobial susceptibility tests for bacteria that grow aerobically; approved standard. M7-A7. Clinical and Laboratory Standards Institute, Wayne, PA, 2006. 63. Osoagbaka, O.U., Evidence for the pathogenic role of Rhodococcus species in pulmonary disease, J. Appl. Bacteriol., 66, 497, 1989. 64. Steingrube, V.A. et al., Rapid identification of clinically signifi­ cant species and taxa of aerobic actinomycetes, including Acti­ nomadura, Gordona, Nocardia, Rhodococcus, Streptomyces, and Tsukamurella isolates, by DNA amplification and restriction endonuclease analysis, J. Clin. Microbiol., 35, 817, 1997. 65. Buchman, A.L. et al., Central venous catheter sepsis caused by unusual Gordona (Rhodococcus) species: identification with a digoxigenin-labeled rDNA probe, Clin. Infect. Dis., 15, 694, 1992. 66. Lasker, B.A. et al., Identification and epidemiological typing of clinical and environmental isolates of the genus Rhodococcus with use of a digoxigenin-labeled rDNA gene probe, Clin. Infect. Dis., 15, 223, 1992. 67. Riegel, P. et al., Rhodococcus chubuensis Tsukamura 1982 is a later subjective synonym of Gordona sputi [Tsukakmura 1978] Stackebrandt 1989 comb.nov., Int. J. Syst. Bacteriol., 44, 764, 1994. 68. Shen, F-T., and Young, C-C., Rapid detection and identification of the metabolically diverse genus Gordonia by 16S rRNA-gene-targeted genus-specific primers, FEMS Microbiol. Lett., 250, 221, 2005. 69. Shen, F-T. et al., Detection of filamentous genus Gordonia in foam sample using genus-specific primers combined with PCR—denaturing gradient gel electrophoresis analysis, Can. J. Microbiol., 53, 768, 2007. 70. Shen, F-T. et al., Molecular detection and phylogenetic characterization of Gordonia species in heavily oil-contaminated soils, Res. Microbiol., 159, 522, 2008. 71. Ruimy, R. et al., A phylogeny of the genus Nocardia deduced from the analysis of small-subunit ribosomal DNA sequences, including transfer of Nocardia amarae to the genus Gordona and Gordona amarae comb. nov., FEMS Microbiol. Lett., 123, 261, 1994. 72. Patel, J.B. et al., Sequence-based identification of aerobic actinomycetes, J. Clin. Microbiol., 42, 2530, 2004. 73. Cloud, J.L. et al., Evaluation of partial 16S ribosomal DNA sequencing for identification of Nocardia species by using the MicroSeq 500 system with an expanded database, J. Clin. Microbiol., 42, 578, 2004. 74. Shen, F-T. et al., Phylogenetic analysis of members of the metabolically diverse genus Gordonia based on proteins encoding the gyrB gene, Res. Microbiol., 157, 367, 2006. 75. Kang, Y. et al., Phylogenetic studies of Gordonia species based on gyrB and secA1 gene analysis, Mycopathologia, 167, 95, 2009. 76. Blanc, V. et al., Gordonia terrae: a difficult-to-diagnose emerging pathogen? J. Clin. Microbiol., 45, 1076, 2007. 77. Kim, B.C., Park, J.H., and Gu, M.B., Multiple and simultaneous detection of specific bacteria in enriched bacterial communities using a DNA microarray chip with randomly generated genomic DNA probes, Anal. Biochem., 77, 2311, 2005. 78. Kim, B.C., Park, J.H., and Gu, M.B., Multiple and simultaneous detection of specific bacteria in enriched bacterial communities using a DNA microarray chip with randomly generated genomic DNA probes, Anal. Biochem., 77, 2311, 2005.

110 79. De los Reyes, F.L. III. et al., Characterization of filamentous foaming in activated sludge systems using oligonucleotide hybridization probes and antibody probes, Water Sci. Tech., 37, 485, 1998. 80. Carr, E.L. et al., Improved permeabilization protocols for fluorescence in situ hybridization (FISH) of mycolic-acid-containing bacteria found in foams, J. Microbiol. Methods, 61, 47, 2005. 81. Santos, S.R., and Ochman, H., Identification and phylogenetic sorting of bacterial lineages with universally conserved genes and proteins, Environ. Microbiol., 6, 754, 2004.

Molecular Detection of Human Bacterial Pathogens 82. Zeigler, D.R., Gene sequences useful for predicting relatedness of whole genomes in bacteria, Int. J. Syst. Evol. Microbiol., 53, 1893, 2003. 83. Heller, L.C., Jones, M., and Widen, R.H., Comparison of DNA pyrosequencing with alternative methods for identification of mycobacteria, J. Clin. Microbiol., 46, 2092, 2008. 84. Ahmadian, A., Ehn, M., and Hober, S., Pyrosequencing: history, biochemistry and future, Clin. Chim. Acta., 363, 83, 2006.

10 Micrococcus and Kocuria Dongyou Liu CONTENTS 10.1 Introduction.......................................................................................................................................................................111 10.1.1 Classification and Biology.....................................................................................................................................111 10.1.2 Morphology and Biochemical Properties..............................................................................................................111 10.1.3 Clinical Features and Pathogenesis.......................................................................................................................113 10.1.4 Diagnosis...............................................................................................................................................................113 10.2 Methods.............................................................................................................................................................................114 10.2.1 Sample Preparation................................................................................................................................................114 10.2.2 Detection Procedures.............................................................................................................................................114 10.3 Conclusion.........................................................................................................................................................................114 References...................................................................................................................................................................................114

10.1  INTRODUCTION 10.1.1  Classification and Biology The genera Micrococcus and Kocuria belong to the family Micrococcaceae, order Actinomycetales, class Actinobacteria, phylum Actinobacteria. Other recognized genera in the family Micrococcaceae include Acaricomes, Arthrobacter, Citricoccus, Nesterenkonia, Renibacterium, Rothia, Sinomonas, Stomatococcus, and Zhihengliuella. The genus Micrococcus, first described in 1872, once contained a diverse group of bacteria that was later dissected into the genera Kocuria, Micrococcus, Nesterenkonia, Kytococcus, and Dermacoccus.1,2 At the present, the genus consists of six species: Micrococcus antarcticus, Micrococcus endophyticus, Micrococcus flavus, Micrococcus luteus, Micrococcus lylae, and Micrococcus yunnanensis.1–7 The species Micrococcus luteus is subdivided into three biovars on the basis of their distinct chemotaxonomic and biochemical traits: biovar I (represented by strain DSM 20030T); biovar II (represented by strains D7, 3, 6, 7, 13C2, 38, 83, and 118); and biovar III (represented by strain Ballarat = DSM 14235 = CCM 4960).3 The genus Kocuria was created in 1995 to cover a number of gram-positive, catalase positive, coagulase negative, strictly aerobic cocci that were classified previously within the genus Micrococcus.2 Kocuria spp. are clearly separable from Micrococcus and other related gram-positive cocci (e.g., Arthrobacter and Rothia) on the basis of distinct 16S rRNA gene sequences.2 As it currently stands, the genus Kocuria (named after Miroslav Kocur) is composed of 16  species: Kocuria aegyptia, Kocuria atrinae, Kocuria carniphila, Kocuria flava, Kocuria gwangalliensis, Kocuria

halotolerans, Kocuria himachalensis, Kocuria koreensis, Kocuria kristinae, Kocuria marina, Kocuria palustris, Kocuria polaris, Kocuria rhizophila, Kocuria rosea, Kocuria salsicia, Kocuria turfanensis, and Kocuria varians.8–18 Members of the genera Micrococcus and Kocuria are distributed in a wide variety of natural environments such as soil, the rhizosphere, freshwater, marine sediments, and fermented foods. In fact, Micrococcus and Kocuria spp. are often used as starter cultures in the manufacture of fermented fish and dry sausages to promote the desired red color development and stability (through reduction of nitrate to nitrite), to prevent rancidity (through peroxide decomposition), to maintain hygienic safety (through nitrite permanence), and to contribute to the flavor of fermented meat products (through formation of esters and other aromatic compounds from amino acids). In addition, Micrococcus and Kocuria spp. are also found in the skin, mucosae, and oropharynx of humans as commensal microflora, and several of them have been associated with human diseases.

10.1.2  Morphology and Biochemical Properties Members of the genus Micrococcus are gram-positive, nonmotile, nonendospore forming, aerobic, and chemoorgano­ trophic cocci. Micrococcus cells are catalase and oxidase positive, mesophilic, and nonhalophilic. The peptidoglycan of Micrococcus spp. contains amino acid l-lysine, which is characteristic of the genus. The predominant menaquinones in Micrococcus cells are either MK-8 and MK-8(H2), although minor amounts of MK-8(H), MK-7 or MK-7(H2), and MK-9(H2) may be present. Micrococcus cells lack mycolic acids and teichoic acids, but may contain teichuronic 111

112

acids as an amino sugar in the cell wall polysaccharide. The G + C content of Micrococcus DNA is 69–76 mol%. Micrococcus spp. are primarily found in mammalian skin. The type species of the genus is Micrococcus luteus.1–3 Members of the genus Kocuria are gram-positive, nonencapsulated, nonendospore forming, chemoorganotrophic, and strictly aerobic cocci. Kocuria spp. are catalase positive, nonhalophilic, and mesophilic. They usually grow on simple media, and are characterized by the presence of menaquinones MK-7(H2) and MK-8(H2), lysine-based A3α-type peptidoglycan, PI-type phospholipids, and saturated branched fatty acids in their cell envelopes.2 The G + C content of Kocuria DNA ranges from 66 to 75 mol%. The type species is Kocuria rosea.10 Among the pathogenic Kocuria spp., Kocuria rosea (rosea, rose colored; formerly Micrococcus rosea) cells are spherical (with a diameter of 1–1.5 µm) and occur in pairs, tetrads, and clusters. Colonies are circular, slightly convex, smooth (occasionally rough), and pink or red, and colony morphology and color tend to become more distinct with age. K. rosea grows well at temperature between 25°C and 37°C and in the presence of NaCl concentrations of up to 7.5% NaC1. Its growth is stimulated by cysteine and thiamine or cysteine, thiamine, and pantothenic acid. In nutrient broth, K. rosea demonstrates moderate turbidity and a deposit. Most K. rosea do not grow on inorganic nitrogen agar and on Simmons citrate agar. K. rosea is oxidase test negative (rarely weakly positive) and benzidine test positive. The bacterium can reduce nitrate, but not nitrite. In addition, it is negative for arginine dihydrolase, phenylalanine deaminase, urease, lecithinase, phosphatase, p-galactosidase, and coagulase. Furthermore, K. rosea does not hydrolyse esculin and gelatin, and its hydrolysis of Tween 80 and starch is variable. Many K. rosea strains can utilize rhamnose, xylose, and glucitol, but form no acid from mannose, galactose, lactose, and glycerol. The G + C content of K. rosea DNA is 66–75 mol%. K. rosea is saprophytic, nonhemolytic, and is often isolated from soil and water, and also occasionally from human infections.2 Kocuria varians (varians, varying; formerly Micrococcus varians) cells are spherical (with a diameter of 0.9–1.5 µm) and occur in tetrads and irregular clusters of tetrads or, rarely, in packets or as single cells. The optimal growth temperature of K. varians ranges from 22°C to 37°C. Colonies (up to 4 mm in diameter) are circular, slightly convex, smooth and glistening, and yellow in color. Some K. varians strains may form rough, wrinkled, matt, dry colonies. Colony morphology and color often become more distinct with age. The bacterium usually grows on Simmons citrate agar but not on inorganic nitrogen agar. In nutrient broth, K. varians shows slight turbidity and a deposit. It can grow on nutrient agar containing up to 7.5%–10% NaCl, and its growth is stimulated by methionine or cysteine and thiamine. K. varians is oxidase test negative; benzidine test positive, in addition to being usually positive for urease and nitrate and nitrite reductases. It is negative for arginine dihydrolase, coagulase, p-galactosidase, phosphatase, lecithinase, phenylalanine deaminase, and DNase. K. varians does not hydrolyse esculin

Molecular Detection of Human Bacterial Pathogens

and Tween 80, and may show varied hydrolysis of starch and gelatin. In addition, it is indole and hydrogen sulfide negative. While the bacterium produces acid from glucose, xylose, and fructose, it does not do so from mannose, maltose, arabinose, raffinose, galactose, rhamnose, ribitol, lactose, mannitol, glycerol, glucitol, inositol, and salicin. The G + C content of K. varians DNA is 66–72 mol%. The bacterium is saprophytic, nonhemolytic, and isolated from mammalian skin, soil, and water.2 Kocuria kristinae (kristinae, of Kristin; formerly Micrococcus kristinae) cells are 0.7–1.1 µm in diameter and occur in tetrads or clusters. The optimal growth temperature of K. kristinae ranges from 25°C to 37°C. Colonies are circular, entire or crenate, convex, smooth or rough, up to 2 mm in diameter, and pale cream to pale orange in color, which becomes more intense with time. K. kristinae is lightly facultatively anaerobic, and grows well on nutrient agar containing up to 10% NaCl, but very weakly in the presence of 15% NaC1. Its growth is stimulated by leucine, lysine, valine, tyrosine, and niacin. K. kristinae is unable to grow on Simmons citrate agar, and rarely grows on inorganic nitrogen agar. Being oxidase and benzidine test positive, the bacterium is negative for arginine dihydrolase, lecithinase, and phosphatase activity, and variable for urease P-galactosidase activity. Most K. kristinae strains do not reduce nitrate. It is positive for hydrolysis of esculin but negative for hydrolysis of gelatin, Tween 80, and starch. It can produce acid aerobically from glucose, fructose, mannose, sucrose, and glycerol, and usually from maltose and glucitol, although it cannot utilize rhamnose, xylose, ribose, arabinose, raffinose, melibiose, mannitol, ribitol, and galactitol; usually does not utilize galactose and lactose. The G + C content of K. kristinae DNA is 67 mol%. It is nonhemolytic, and its primary habitat is human skin. K. kristinae has been associated with catheter-related bacteremia.2 Kocuria rhizophila (rhiza, a root; philos, loving; rhizophila root-loving) cells are gram-positive, nonacid-fast, nonmotile, nonendospore forming, aerobic, chemoorganotrophic, and spherical with a diameter of 1.0–1.5 µm, and occur in pairs, tetrads, and packets. Colonies are 1.5– 2.5 mm in diameter, and are opaque, smooth with irregular edges with yellow pigmentation. The bacterium grows well between pH 5.7–7.7, in the presence of 10% NaCl, and on inorganic nitrogen and Simmons’ citrate, but does not grow at temperature above 40°C. It is positive for catalase, benzidine gelatinase, phosphatase, and Tween 80 hydrolysis, but negative for oxidase, indol, urease, arginine dihydrolase and phenylalanine deaminase, and esculin and starch hydrolysis. The bacterium does not reduce nitrate to nitrite. It produces acid production from d-glucose, d-fructose, d-mannose, and saccharose, and also utilizes adonitol, l-arabinose, d-fructose, l-fucose, d-glucose, turanose, xylitol, methylpyruvate, glucuronamide, dextrin, glycogen, Twee 40, Tween 80, and N-acetyl-d-glucosamine. The major menaquinones of K. rhizophila are MK-7(H2) and MK-8(H2). The genome of K. rhizophila DC2201 (NBRC 103217) has been shown to harbor a single circular chromosome of 2,697,540 bp in

113

Micrococcus and Kocuria

length with an average G + C content of 71.16%. Of the 2357 predicted protein-coding genes, 87.7% encode proteins that are orthologous to actinobacterial proteins.19 K. rhizophila was first isolated from the rhizosphere of Typha angustifolia from a floating mat on the Soroksar tributary of the river Danube, Hungary8 and a human case of K. rhizophila infection was documented recently. Kocuria marina (marina, of the sea) cells are grampositive, aerobic, nonmotile, halotolerant cocci that grow at 4°C–43°C. The bacterium is positive for catalase, β-galactosidase, and urease, but negative for arginine dihydrolase, lysine and ornithine decarboxylase, oxidase, and alkaline phosphatase. It reduces nitrate, but does not produce hydrogen sulfide. In addition, it produces acid from l-fucose, although it is negative for indole and acetoin (Voges–Proskauer reaction). It can hydrolyse casein, gelatin, and Tween 40, but not agar, alginate, cellulose, DNA, starch, Tween 20, and Tween 80. Furthermore, it utilizes glucose, lactose, mannose, and sucrose as sole carbon sources, but not adonitol, l-arabinose, meso-inositol, mannitol, sorbitol, citric acid, or malonic acid. The G + C content of K. marina DNA is 60 mol%.11 The bacterium was first isolated from marine environment, but a case of human infection with this bacterium was recently reported.11

10.1.3  Clinical Features and Pathogenesis Micrococcus and Kocuria spp. are widespread in nature and are frequently found as normal skin flora in humans and other mammals. Micrococcus strains such as M. luteus have been associated with catheter-related bacteremia in patients undergoing hemodialysis or leukaemia treatment, pneumonia, endocarditis, intracranial abscesses, continuous ambulatory dialysis peritonitis, septic arthritis, and meningitis.20–27 Since Kocuria spp. were contained in the genus Micrococcus prior to 1995, some of the early reports documenting the roles of Micrococcus strains in human infections may be attributable to Kocuria spp. In recent years, a number of Kocuria-related infections have been documented. Basaglia et al.28 described the first case of a catheter-related recurrent bacteremia due to Kocuria kristinae in a 51-year-old woman with a poorly differentiated ovarian carcinoma. The patient had a permanent central venous catheter (CVC) implanted for chemotherapy and developed neutropenia and sepsis a few months afterwards. Kocuria kristinae isolates grew from three blood samples cultured aerobically onto sheep blood agar plates at 37°C for 18 h. They were nonhemolytic, catalase positive, strictly aerobic, nonmotile, and unable to reduce nitrate to nitrites or to hydrolyze gelatin, arginine, and esculin. The bacterial isolates produced acid from trehalose, glucose, fructose, mannose, glycerol, and saccharose but not from mannitol, raffinose, arabinose, lactose, or ribose. This identification was confirmed with the commercially available ID32 Staph ATB system (Biomerieux), as well as sequencing analysis of the partial 16S rRNA gene (rDNA; 642 bp). The patient recovered after removal of CVC and antibiotic treatment.

Altuntas et al.29 reported the first case of a catheter-related bacteremia caused by Kocuria rosea in a 39-year-old man undergoing peripheral blood stem cell transplantation due to relapsed Hodgkin disease. The patient presented chills, high fever (temperature 38.5°C), tachycardia (heart rate 108 beats/ min), and increased respiratory rate 3 days after implantation of a Hickman-type central venous catheter. Serial blood cultures taken from the catheter and a peripheral vein revealed the presence of Kocuria rosea as confirmed with the commercial tests (ID32 Staph ATB system, and mini-API, Biomerieux). The patient recovered after treatment with vancomycin and by catheter removal. Ma et al.30 described the first case of K. kristinae-related acute cholecystitis in a patient who underwent laparoscopic cholecystectomy. The patient developed postoperative fever and recovered after levofloxacin treatment. Becker et al.31 documented the first case of a Kocuria rhizophila infection in a boy with methylmalonic aciduria due to a noncobalaminresponsive deficiency of methylmalonyl coenzyme A mutase. The patient developed sepsis two years after a subcutaneous implantable vascular-access port (Port-A-Cath; Vital-Port) was placed in his left internal jugular vein. The patient showed the signs of acute pancreatitis and fever. A Kocuria rhizophila strain was isolated from blood samples drawn through a port system and from peripheral veins during septic episodes. The bacterium resembles Micrococcus species as assessed by the Bactec 9240 system (Becton Dickinson). It is gram-positive, catalase positive and oxidase negative, and appears in pairs, tetrads, and packets. Colonies are smooth and circular with a yellow tinge and appeared dull and creamy on Mueller-Hinton blood agar under strictly aerobic conditions. 16S rRNA gene sequence analysis of the bacterium confirmed its K. rhizophila identity. Lee et al.32 reported the first two cases of Kocuria marina peritonitis in patients undergoing continuous ambulatory peritoneal dialysis. Furthermore, Lai et al.33 isolated Kocuria bacteria in blood specimens from five immunocompromised patients. Of these patients, three had catheter-related bacteremia and one had infective endocarditis caused by K. kristinae, and one had a K. marina isolate.

10.1.4  Diagnosis Micrococcus and Kocuria spp. are cultured on sheep blood agar, MacConkey agar, and chocolate agar at 35°C for 48 h. Gram-positive cocci from pale cream colonies after two days incubation appear in tetrads. The organisms are nonhemolytic, catalase positive, coagulase negative, and nonmotile, and can be characterized by using bioMérieux ID32 Staph ATB system, BD Phoenix PMC/ID-13 system (Becton Dickinson), and Vitek 2 (bioMérieux) as well as by their antibiotic profiles.34 Further confirmation is often achieved by using molecular techniques. Sequencing analysis of the partial or full 16S rRNA gene has proven valuable. Other molecular techniques for Micrococcus and Kocuria spp. include ribotyping (e.g., automated ribotyping with EcoRI and PvuII as restriction enzymes based on the RiboPrinter® Microbial

114

Characterization System (DuPont-Qualicon, Wilmington, DE), denatured gradient gel electrophoresis (DGGE) analy­ sis of PCR amplified 16S rDNA V3 region, the 16S-23S rRNA intergenic spacer region (ISR)-PCR, random amplified polymorphic DNA (RAPD), pulsed-field gel electrophoresis (PFGE), and rpoB gene sequencing.35

10.2  METHODS

Molecular Detection of Human Bacterial Pathogens





10.2.1  Sample Preparation BacT/Alert blood-culture bottles are inoculated with 8–10 mL blood and incubated for 7 days. Gram-positive isolates from primary cultures are grown on 5% blood agar and MacConkey agar for 48 h at 37°C under aerobic conditions.36 The blood bottles are kept at –20°C until the DNA extraction is performed. Bacterial DNA is extracted from the bottles by the alkali wash/lysis method. Extracted DNA is stored at –20°C prior to PCR.36

10.2.2  Detection Procedures (i) Sequencing analysis of complete 16S rRNA gene  Paster et al.37 described the application of universal primers 27F (5′-AGAGTTTGATCCTGGCTCAG-3′, corresponding to positions 8–27 in Escherichia coli 16S rRNA gene) and 1492R (5′-TACCTTGTTACGACTT-3′, corresponding to positions 1492–1507) for amplification of nearly the entire length of the 16S rRNA gene from bacteria including Kocuria and Micrococcus spp. Procedure 1. PCR mixture (100 µL) is made up of 50 mM KCl, 10 mM Tris pH 8.3, 1.5 mM MgCl2, 2 mM each of dNTP, 1 µM (each) primers 37F and 1492R, 2.5 U of Taq DNA polymerase, and 1 pg to 100 ng of template DNA. 2. The mixture is subjected to an initial denaturation at 94°C for 5 min, 25 cycles of 94°C for 60 s, 50°C for 30 s, and 72°C for 2 min, and a final extension step at 72°C for 5 min. 3. The resulting PCR products are then sequenced by the use of ABI technology with the same universal primers. The nucleotide sequence is analyzed with BLAST program. (ii) Sequencing analysis of partial 16S rRNA gene  Karahan et al.36 utilized primers fD1 (5′ AGAGTT TGATCCTGGCTCAG 3′) and 800R (5′ GAGTACC AGGGTATCTAATCC 3′) to amplify an 800 bp region of the gene encoding the 16S rRNA of eubacteria. Procedure 1. PCR mixture (50 µL) is made up of 5 µL extracted bacterial DNA, 25 pmol of each primer, 200 µmol each dNTP, 1.5 mM MgCl2 and 1.5 U Taq DNA polymerase (Fermentas) and appropriate 1 × PCR buffer.



2. PCR amplification is conducted with an initial denaturation at 94°C for 5 min, 35 cycles of 94°C for 1 min, 60/58/55°C for 1 min (2/2/31 cycles), and 72°C for 1 min, and a final extension at 72°C for 4 min. 3. A portion (10 µL) of the PCR products are subjected to 2% (w/v) agarose gel electrophoresis, stained with ethidium bromide, and visualized under UV. 4. The remaining PCR products (40 µL) are purified by the EZNA Cycle-Pure Kit (Omega Bio-tek), and subjected to automated sequencing with a CEQ2000XL sequencer (Beckman Coulter) by using the fD1 and 610R (5′-TACCGCGGCTGCTGGCAC-3′) primers. 5. The resulting nucleotide sequences are matched to existing GenBank sequences by nucleotide-nucleotide BLAST analysis.

Alternatively, the 16S and 23S rRNA intergenic region may be amplified and sequenced. Another useful target is the housekeeping gene rpoB, which harbors sufficient nucleotide differences for discrimination among members of Micrococcus and Kocuria spp.35

10.3  CONCLUSION Members of the genera Micrococcus and Kocuria are grampositive, catalase positive, coagulase negative, strictly aerobic cocci that are found ubiquitously in the environments and are present in the skin, mucosae, and oropharynx of humans and animals as commensal microflora. Several Micrococcus and Kocuria species are opportunistic human pathogens that have been associated with catheter-related bacteremia, pneumonia, intracranial abscesses, septic arthritis, meningitis, peritonitis, and endocarditis in human patients, particularly immunocompromised individuals. As Micrococcus and Kocuria species demonstrate phenotypic variability, they are difficult to identify with standard culture and biochemical tests,38 and the true incidence of Micrococcus and Kocuria infections may have been underestimated. Recent development and application of PCR and DNA sequencing techniques have facilitated rapid, precise identification and determination of these bacteria. Consequently, an increasing number of clinical cases due to Micrococcus and Kocuria species have been reported. This highlights the value of molecular tests in the clinical diagnosis of hard-to-identify organisms, such as Micrococcus and Kocuria spp.

REFERENCES



1. Kloos, W.E., Tornabene, T.G., and Schleifer, K.H., Isolation and characterization of micrococci from human skin, including two new species: Micrococcus lylae and Micrococcus kristinae, Int. J. Syst. Bacteriol., 24, 79, 1974. 2. Stackebrandt, E. et al., Taxonomic dissection of the genus Micrococcus: Kocuria gen. nov., Nesterenkonia gen. nov., Kytococcus gen. nov., Dermacoccus gen. nov., and Micrococcus Cohn 1872 gen. emend., Int. J. Syst. Bacteriol., 45, 682, 1995.

Micrococcus and Kocuria







3. Wieser, M. et al., Emended descriptions of the genus Micrococcus, Micrococcus luteus (Cohn 1872) and Micrococcus lylae (Kloos et al. 1974), Int. J. Syst. Evol. Microbiol., 52, 629, 2002. 4. Liu, H. et al., Characterization of Micrococcus antarcticus sp. nov., a psychrophilic bacterium from Antarctica, Int. J. Syst. Evol. Microbiol., 50, 715, 2000. 5. Liu, X.Y. et al., Micrococcus flavus sp. nov., isolated from activated sludge in a bioreactor, Int. J. Syst. Evol. Microbiol., 57, 66, 2007. 6. Chen, H.H. et al., Micrococcus endophyticus sp. nov., isolated from surface-sterilized Aquilaria sinensis roots, Int. J. Syst. Evol. Microbiol., 59, 1070, 2009. 7. Zhao, G.Z. et al., Micrococcus yunnanensis sp. nov., a novel actinobacterium isolated from surface-sterilized Polyspora axillaris roots, Int. J. Syst. Evol. Microbiol., 59, 2383, 2009. 8. Kovacs, G. et al., Kocuria palustris sp. nov. and Kocuria rhizophila sp. nov., isolated from the rhizoplane of the narrowleaved cattail (Typha angustifolia), Int. J. Syst. Bacteriol., 49, 167, 1999. 9. Tang, J.S., and Gillevet, P.M., Reclassification of ATCC 9341 from Micrococcus luteus to Kocuria rhizophila, Int. J. Syst. Evol. Microbiol., 53, 995, 2003. 10. Reddy, G.S. et al., Kocuria polaris sp. nov., an orangepigmented psychrophilic bacterium isolated from an Anta­ rctic cyanobacterial mat sample, Int. J. Syst. Evol. Microbiol., 53, 183, 2003. 11. Kim, S.B. et al., Kocuria marina sp. nov., a novel actinobacterium isolated from marine sediment, Int. J. Syst. Evol. Microbiol., 54, 1617, 2004. 12. Tvrzova, L. et al., Reclassification of strain CCM 132, previously classified as Kocuria varians, as Kocuria carniphila sp. nov., Int. J. Syst. Evol. Microbiol., 55, 139, 2005. 13. Li, W.J. et al., Kocuria aegyptia sp. nov., a novel actinobacterium isolated from a saline, alkaline desert soil in Egypt, Int. J. Syst. Evol. Microbiol., 56, 733, 2006. 14. Mayilraj, S. et al., Kocuria himachalensis sp. nov., an actinobacterium isolated from the Indian Himalayas, Int. J. Syst. Evol. Microbiol., 56, 1971, 2006. 15. Zhou, G. et al., Kocuria flava sp. nov. and Kocuria turfanensis sp. nov., airborne actinobacteria isolated from Xinjiang, China, Int. J. Syst. Evol. Microbiol., 58, 1304, 2008. 16. Park, E.J. et al., Kocuria atrinae sp. nov., isolated from traditional Korean fermented seafood, Int. J. Syst. Evol. Microbiol., 60, 914, 2010. 17. Seo, Y.B. et al., Kocuria gwangalliensis sp. nov., an actinobacterium isolated from seawater, Int. J. Syst. Evol. Microbiol., 59, 2769, 2009. 18. Yun, J.H. et al., Kocuria salsicia sp. nov., isolated from salt-fermented seafood, Int. J. Syst. Evol. Microbiol., 2010 March 12. 19. Takarada, H. et al., Complete genome sequence of the soil Actinomycete Kocuria rhizophila, J. Bacteriol., 190, 4139, 2008.

115 20. Souhami, L. et al., Micrococcus luteus pneumonia: A case report and review of the literature, Med. Pediatr. Oncol., 7, 309, 1979. 21. Fosse, T. et al., Meningitis due to Micrococcus luteus, Infection, 13, 280, 1985. 22. Wharton, M. et al., Septic arthritis due to Micrococcus luteus, J. Rheumatol., 13, 659, 1986. 23. Selladurai, B. et al., Intracranial suppuration caused by Micrococcus luteus, Br. J. Neurosurg., 7, 205, 1993. 24. von Eiff, C. et al., Micrococcus luteus as a cause of recurrent bacteremia, Pediatr. Infect. Dis. J., 15, 711, 1996. 25. Peces, R. et al., Relapsing bacteraemia due to Micrococcus luteus in a haemodialysis patient with a Perm-Cath cathether, Nephrol. Dial. Transplant., 12, 2428, 1997. 26. Shanks, D. et al., Fatal Micrococcus sp. infection in a child with leukaemia—a cautionary case, Med. Pediatr. Oncol., 37, 553, 2001. 27. Ramos, E.R. et al., The crucial role of catheters in micrococcal bloodstream infections in cancer patients, Infect. Control Hosp. Epidemiol., 30, 83, 2009. 28. Basaglia, G. et al., Catheter-related bacteremia due to Kocuria kristinae in a patient with ovarian cancer, J. Clin. Microbiol., 40, 311, 2002. 29. Altuntas, F. et al., Catheter-related bacteremia due to Kocuria rosea in a patient undergoing peripheral blood stem cell transplantation, BMC Infect. Dis., 4, 62, 2004. 30. Ma, E.S. et al., Kocuria kristinae infection associated with acute cholecystitis, BMC Infect. Dis., 5, 60, 2005. 31. Becker, K. et al., Kocuria rhizophila adds to the emerging spectrum of micrococcal species involved in human infections, J. Clin. Microbiol., 46. 3537, 2008. 32. Lee, J.Y. et al., Two cases of peritonitis caused by Kocuria marina in patients undergoing continuous ambulatory peritoneal dialysis, J. Clin. Microbiol., 47, 3376, 2009. 33. Lai, C.C. et al., Catheter-related bacteremia and infective endocarditis caused by Kocuria species, Clin. Microbiol. Infect., 2010 [Epub ahead of print]. 34. Szczerba, I., Susceptibility to antibiotics of bacteria from genera Micrococcus, Kocuria, Nesternkonia, Kytococcus and Dermacoccus, Med. Dosw. Mikrobiol., 55, 75, 2003. 35. Hellmark, B., Söderquist, B., and Unemo, M., Simultaneous species identification and detection of rifampicin resistance in staphylococci by sequencing of the rpoB gene, Eur. J. Clin. Microbiol. Infect. Dis., 28, 183, 2009. 36. Karahan, Z.C. et al., PCR evaluation of false-positive signals from two automated blood-culture systems, J. Med. Microbiol., 55, 53, 2006. 37. Paster, B.J. et al., Prevalent bacterial species and novel phylotypes in advanced noma lesions, J. Clin. Microbiol., 40, 2187, 2002. 38. Ben-Ami, R. et al., Erroneous reporting of coagulase-negative staphylococci as Kocuria spp. by the Vitek 2 system, J. Clin. Microbiol., 43, 1448, 2005.

11 Mycobacterium Shubhada Shenai and Camilla Rodrigues CONTENTS 11.1 Introduction.......................................................................................................................................................................117 11.1.1 Classification, Morphology, and Epidemiology.....................................................................................................117 11.1.2 Clinical Features and Pathogenesis.......................................................................................................................118 11.1.3 Diagnosis...............................................................................................................................................................119 11.1.3.1 Phenotypic Techniques...........................................................................................................................119 11.1.3.2 Molecular Biological Techniques.......................................................................................................... 120 11.2 Methods............................................................................................................................................................................ 122 11.2.1 Sample Preparation............................................................................................................................................... 122 11.2.2 Detection Procedures............................................................................................................................................ 122 11.3 Conclusion and Future Perspectives................................................................................................................................. 124 References.................................................................................................................................................................................. 124

11.1  INTRODUCTION The genus mycobacterium comprises more than 125 species, some of which are pathogenic or potentially pathogenic to human and animals and some of which are saprobes. A variety of diseases, including pulmonary disease, skin and soft tissue infections, lymphadenitis, and disseminated disease, are caused by mycobacteria. The most important pathogen of this genus is M. tuberculosis (MTB) complex, the causative agent of tuberculosis (TB).1 Current global estimates indicate that about one-third of the world’s population is infected with MTB; 8.8 million individuals develop the disease annually. In addition, there are increasing reports of nontuberculous mycobacteria (NTM) infections not only in the immunocompromised but also in immunocompetent hosts.2,3 NTM may cause both asymptomatic infection and symptomatic disease in humans. They infect many sites within the body but primarily cause pulmonary disease, cervical lymphadenopathy, and localized skin and soft tissue lesions.

11.1.1  Classification, Morphology, and Epidemiology Mycobacteria are believed to be among the oldest bacteria on earth, and are ubiquitous in the environment (Table 11.1). Mycobacteria are slender rods that sometimes show branching filamentous forms resembling fungal mycelium, hence the name “mycobacteria,” meaning fungus like. (The Latin prefix “myco” means both fungus and wax; its use here relates to the “waxy” compounds in the cell wall.)

All the members of this genus are aerobic nonspore forming, noncapsulated, nonmotile, single cell bacteria and cell shape varies from that of coccobacilli to elongated rods, approximately 0.2–0.8 µm wide by 1–10 µm long. They are classified as gram-positive bacteria but most of them do not stain well with Gram-stain.4 The unique character of mycobacteria is that they are acid-fast. Acid-fastness is essentially due to the uniquely thick cell wall, which is composed of an interlacing layer of lipids, peptidoglycans, and arabinomannans.4,5 Many mycobacterium species adapt readily to growth on simple substrates, using ammonia or amino acids as nitrogen sources and glycerol as a carbon source in the presence of mineral salts. Optimum growth temperatures vary widely according to the species and range from 23°C to over 50°C. Some species can be difficult to culture and are fastidious in their growth requirements, sometimes taking over two years to cultivate in culture.6 Furthermore, some species also have extremely long reproductive cycles—M. leprae, may take more than 20 days for one division cycle.1 In 1954, Timpe and Runyon classified all mycobacteria into four groups on the basis of growth rate, pigmentation, colonial morphology, and biochemical test characterization. Organisms that took less than 7 days to grow isolated colonies on solid media were called rapidly growing mycobacteria (RGM) and those that took more than 7 days were called as slowly growing mycobacteria (SGM). MTB is an obligate pathogen of humans and is rarely identified in other mammals. It is transmitted from person to person, and it has no significant environmental reservoirs. A number of clinically important NTM are found in soil, water, 117

118

Molecular Detection of Human Bacterial Pathogens

TABLE 11.1 Scientific Classification Kingdom

Bacteria

Phylum

Actinobacteria

Order Suborder Family

Actinomycetales Corynebacterineae Mycobacteriaceae

Genus Clinically relevant species

Mycobacterium M. tuberculosis complex, M. leprae, M. avium, M. intracellulare, M. kansasii, M. marinum, M. ulcerans, M. simiae, M. gordonae, M. genavense, M. szulgai, M. scrofulaceum, M. haemophilum, M. fortuitum complex, M. chelonae, M. abscessus, etc.

domestic and wild animals, milk and other food materials. They may contaminate clinical specimens from the environment or transiently colonize body surfaces and sections. Most infections with NTM appear to be acquired by aspiration or inoculation of the organisms from the natural reservoir. There is no firm evidence of human-to-human or animal-to-human transmission of NTM disease.7 The geographic distribution of some species does not appear to be uniform. Depending on the locality, infections caused by other mycobacteria have been reported to account for 0.5%–30% of all mycobacterial infections.8–10

11.1.2  Clinical Features and Pathogenesis There are five main types of disease caused by mycobacteria: TB, pulmonary infections resembling TB, localized skin and soft tissue infections, localized lymph node involvement, and disseminated disease. TB. The clinical manifestation and eventual outcome of TB depends on the virulence of the pathogen and the nature of the host’s immune response. When a person inhales air that contains droplets, most of the larger droplets are lodged in the upper respiratory tract (i.e., nose and throat), where infection is unlikely to develop. However, the droplet nuclei may reach the small air sacs of the lung (the alveoli) where infection begins. When organisms reach the lungs, there are four potential outcomes. (i) The initial host response can be completely effective and kill all the organisms; in this case the patient has no chance of developing TB. (ii) The organism can begin to multiply and grow immediately after the infection, causing clinical disease known as primary tuberculosis. (iii) The bacilli may remain dormant and do not cause disease. This is referred to as latent infection indicated only by a positive tuberculin skin test. (iv) There could be reactivation of the latent organisms causing postprimary infection.11 Some bacilli may be killed immediately by the process of phagocytosis, which involves binding of the bacterium to the host cell, internalization, and finally growth inhibition or killing.11 If the bacilli are able to survive the initial defenses,

they multiply within the alveolar macrophages and a small number of bacilli enter the bloodstream and disseminate. Intracellular multiplication of the bacillus is interrupted only with the development of specific cellular immunity, which sets in about 6–8 weeks after infection. Once the immune response called cell-mediated hypersensitivity is established, the T cells usually respond to the tuberculin skin test (PPD test) and produce a characteristic red welt. Some of the T cell signals produce inflammatory reactions; other signals recruit and activate specialized cells to kill bacilli and wall off infected macrophages in tiny, hard capsules known as tubercles. A tubercle consists of a few small lymphocytes and a compact collection of activated macrophages, which sometimes differentiate into epitheloid cells or multinucleated giant cells. The massive activation of macrophages that occurs within tubercles often results in the concentrated release of lytic enzymes.4 These enzymes destroy nearby healthy cells, resulting in a circular region of necrotic tissue, which eventually form a lesion with a caseous consistency. As these caseous lesions heal, they calcify and are visible on X-rays, where they are called Ghon complexes. For majority of individuals with the normal immune function, the initial infection with M. tuberculosis appears to be arrested once cell-mediated immunity develops, although small number of viable bacilli within the granuloma may persist. If the cell-mediated immunity is unable to control the infection, the individual progresses to active disease called progressive primary tuberculosis.12 Individuals infected with MTB have a 10% lifetime risk of developing the disease. The remaining 90% will stay infected, but be free of disease for the rest of their lives.13 Pulmonary Infections Resembling TB. Chronic pulmonary disease is the most common localized clinical manifestation of NTM. MTB complex is the most frequent pathogen causing lung disease, followed by M. avium complex (MAC), M. kansasii, M. abscessus, M. fortuitum, M.  szulgai, M. simiae, M. xenopy, M. malmoense, M. celatum, M. asiaticum, and M. shimodii. Respiratory infections due to NTM are often associated with various conditions such as chronic obstructive pulmonary disease, cystic fibrosis of lung, bronchiectasis, emphysema of lung, previously treated pulmonary TB, and lung cancer. Other predisposing conditions include diabetes mellitus, leukemia, systemic lupus erythematosus, chronic alcoholism, AIDS, and transplant patients.14,15 The interpretation of NTM in the sputum of HIV-positive patients presents a particular problem, as these patients are frequently infected with NTM without evidence of pulmonary disease.15,16 Localized Skin and Soft Tissue Infections. Skin and soft tissue infections in immunocompetent patients are the result of direct exposure to contaminated water sources through inoculation after accidental trauma, surgery, and injection. Watery skin lesions may be caused due to the inoculation of opportunistic mycobacteria into superficial abrasions. Such infections are usually caused by M. marinum and occur in users of swimming pools and keepers of tropical fish, hence

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Mycobacterium

the names “swimming pool granuloma” and “fish fancier’s finger.”17–19 Other species such as M. kansasii, M. abscessus, and M. chelonae cause similar lesions. Infections due to M. ulcerans lead to necrosis of subdermal tissue and secondary skin ulceration. Post injection mycobacterial abscesses are usually due to the rapidly growing species M. fortuitum, M. abscessus, and M. chelonae. These may occur sporadically in diabetics,20 for example, or in “epidemics” when several individuals are injected with a contaminated batch of a vaccine or drug.21 Similar infections, caused by rapid growers, results in penetrating injuries surgery. Serious outbreaks of sternal infections have followed open heart surgery.22 A number of cases of corneal infection by rapid growing species have also occurred, presumably as a result of direct implantation.23 M. haemophilium is a rare cause of nodular or ulcerative skin lesions and all reported infections have occurred in immunosuppressed individuals, particularly recipients of renal transplants.17,24 Localized Lymph Node Involvement. Lymphadeno­ pathy due to NTM is usually cervical, unilateral, and selflimiting. Most cases occur in children under the age of 5 years.25 The most common species found in lymphadenitis in children are M. avium, M. intracellulare, M. scrofulaceum, M. malmoense, M. kansasii, M. haemophilum, and M interjectum.25 In adults, the infection may occur as part of a disseminated infection. Disseminated Disease. This form of nonpulmonary disease is similar to that caused by MTB complex. Thus, single or multiple lesions may occur in bone, urinary tract, central nervous system, lymph nodes and skin. In other cases, the infection resembles cryptic disseminated TB with multi-­organ involvement. In such cases there is little or no histological evidence of an immune response: indeed, the bone marrow may be teeming with acid-fast bacilli while appearing normal when stained with the usual hematological stains.25 Such disseminated disease usually occurs in association with congenital or acquired immunodeficiency affecting cell-mediated immunity, including therapeutic immunosuppresion following transplant surgery26 and AIDS.16 Most cases of disseminated disease are due to MAC or M. chelonae.

11.1.3  Diagnosis Presumptive diagnosis of mycobacterial infection is generally made on the basis of patient’s history, clinical presentation of disease, radiological findings, and presence of acid-fast bacilli (AFB) in the patient’s specimen. The most commonly used methods for diagnosis of mycobacterial diseases are listed in Table 11.2. Clinical symptoms and radiological findings, though useful for preliminary diagnosis, may not always provide absolute prediction of the disease. Serological tests for TB are often inconclusive. Microscopy is rapid, easy to perform, and specific; however, it lacks sensitivity, requiring >10,000 bacilli/mL of sputum to become positive. Further, microscopy cannot distinguish between viable and nonviable cells, between different species of mycobacteria and drugsusceptible or drug-resistant M. tuberculosis complex.27 Accurate, rapid microbiological diagnosis of TB and other mycobacterial infections begin with proper specimen collection and the rapid transport of the specimen to the laboratory. Clearly labeled specimen must be transported to the laboratory quickly because some tests, such as AFB smears and nucleic acid amplification (NAA) procedures, can reliably be reported within 24–48 h of receipt. If processing is delayed, the specimens should be refrigerated properly. As NTMs are ubiquitous in nature and may sometimes be isolated as laboratory contaminants, the isolation of these organisms from specimens should meet certain criteria and guidelines to confirm their etiological significance.14 11.1.3.1  Phenotypic Techniques Culture. Isolation of mycobacteria by culture is sensitive and regarded as the gold standard. It has been estimated to detect 10–100 viable mycobacteria per ml of sample.28 Another advantage of culture is that it allows specific species identification and testing for recognition of drug susceptibility patterns. With the emergence of drug-resistant tuberculosis, accurate, rapid, and cost-effective tests to detect MDRTB and XDRTB presents a challenge to TB control activities. Most laboratories in disease endemic region rely on conventional Lowenstein & Jensen (L.J) media for culture followed by use of different biochemical tests for identification of mycobacteria. These conventional culture and identification

TABLE 11.2 Most Commonly Used Methods for Diagnosis of Mycobacteria Method

Advantages

Disadvantages

Clinical signs and symptoms Radiology (X-ray, CT, MRI) Serology Microscopy Conventional culture (egg/agar based) Automated liquid culture Commercial hybridization probe assay PCR (in-house/kit based) Sequencing

Rapid Readily available Rapid, cost effective Rapid, cost effective Specific, cost effective Specific Rapid Rapid Rapid, gold standard

Not specific, not conclusive, not always present Nonspecific and inconclusive Not sensitive and nonspecific Limited sensitivity and specificity Time consuming (4–10 weeks) Slow (2–3 weeks), expensive Expensive, demanding technical expertise and not available for all species Expensive, demanding technical expertise and quality control at each step Expensive

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tests are complex, laborious, time consuming, and many times fail to provide precise identification. Newer phenotypic techniques aim for a more rapid detection of growth by using metabolic activities of growing bacteria. In contrast, automated liquid culture-based techniques can detect growth dependent changes such as CO2 production [BACTEC 460 (Becton Dickinson, Sparks, MD), MB/BacT (bioMerieux)] or O2 consumption [Mycobacterium Growth Indicator Tube (MGIT (Becton Dickinson, Sparks, MD)) 960, Versa TREK (Trek Diagnostics)] and help in rapid diagnosis of disease. Liquid culture systems, when combined with DNA probes or the Capilia TB assay (TAUNS, Numazu, Japan) for rapid species identification, are capable of producing positive results in 2 weeks for the vast majority of sputum smear-positive specimens, and within 3 weeks for smear-negative specimens.11 Mycobacteriophages. The technology uses bacteriophages to infect live MTB and detect the bacilli using either the phage-amplification (FASTPlaque TBTM) method29–31 or the detection of light (Luciferase reporter phage technique).32 Amplification-based assays are commercially available and have specificity of 98.7%–99% and sensitivity of 70%–75% compared to conventional L.J method.29–31 In general, phage assays have a turn around time of 2–3 days and require a laboratory infrastructure similar to that required for performing mycobacterial cultures. High Performance Liquid Chromatography (HPLC). HPLC is based on the observation that each mycobacterium species synthesizes a unique set of mycolic acids, β-hydroxyα-fatty acids, that are components of the cell wall.33,34 HPLC can produce a pattern that reliably identifies and distinguishes more than 50 Mycobacterium species. It can be performed in a few hours from pure cultures. The initial equipment cost for HPLC limits its availability. Due to various limitations of phenotypic methods and HPLC in recent years, molecular techniques have become increasingly popular, as they are rapid, reliable, highly sensitive, and can be used directly for large number of clinical specimens. 11.1.3.2  Molecular Biological Techniques Molecular methods have several advantages over culturebased techniques and include a shorter turn around time, no requirement for culture of an organism, the possibility of direct application in clinical specimens, less biohazard risk, and feasibility for automation. Nucleic acid amplification tests (NAA), also called direct amplification tests, are available either as commercial kits or also widely used with inhouse assays. COBAS Amplicor. The Amplicor MTB test (Roche Molecular Systems, Basel, Switzerland) relies on the standard polymerase chain reaction (PCR). A 584-bp fragment of the 16S ribosomal RNA gene, comprising a species-specific region flanked by genus-specific sequences, is amplified using biotinylated primers. Then the use of magnetic beads coated with specific probes allows the removal of any other DNA by washing. The detection of the specific amplification product is performed by adding an avidinenzyme conjugate and a chromogenic subject. The system

Molecular Detection of Human Bacterial Pathogens

uses uracil-N-glycosylase in the master mix for contamination control. The enzyme uracil-N-glycosylase, added to the samples before amplification, destroys any amplicons resulting from previous amplifications without damaging the uracil free target DNA. From the literature review, specificity of this USFDA-approved test is close to 100% and sensitivity ranges from 90% to 100% in smear-positive samples and from 50% to 95.9% in smear-negative ones.35,36 BD Probe Tec ET. The BD Probe Tec ET system (Becton Dickinson, Sparks, MD) uses DNA polymerase and isothermal strand displacement amplification to produce multiple copies of IS6110, an insertion element to MTB complex. The turn around time is 3.5–4 h. The literature reports a rate of sensitivity ranging from 98.5% to 100% for smear-positive samples and variable (0.33%–100%) for smear-negative ones.37,38 Inhouse PCR assays are used predominantly in developing countries, where commercial NAA are expensive, and they vary widely in their target gene sequences, protocols, and laboratory methods. The accuracy of inhouse PCR test has been more heterogeneous than commercial kits.39 Inhouse tests are laboratory PCR is highly sensitive (96%) and specific (97%–100%). However, its relationship with the clinical disease needs to be ascertained.40,41 Molecular identification of mycobacteria targeting different genes. like IS6110, 16S rRNA, 65 kiloDalton (kDa) heatshock protein gene (hsp 65), 32-kDa protein genes, the gene encoding 126-kDa fusion protein, RD region, and rpoβ gene have been reported.42–46 The most commonly used mycobacterial DNA sequences for PCR are IS6110 (IS986), 16S rRNA, and the gene encoding 65 kDa. An insertion sequence (IS) designated IS6110 is specific for M. tuberculosis complex and generally occurs in 1–20 copies per cell, making it an ideal target for amplification in inhouse PCR protocols. PCR detection of IS6110 in pulmonary and extrapulmonary TB, when compared to culture, has a sensitivity, specificity, and positive predictability of 83.5%, 99%, and 94.2% respectively.46 PCR Restriction Enzyme Analysis (PCR RE). The PCR RE method is based on the amplification of a fragment of the target gene by PCR followed by the digestion of the amplified product with different restriction enzymes (RE).47–51 The products of the digestion reaction are then separated and visualized on agarose gel electrophoresis. In several recent studies PCR RE has performed well, as compared to the conventional methods.47–51 This simple and cost-effective method has been applied to several genes, such as 16S ribosomal RNA (rRNA), rpoβ, hsp65, gyrB, and dnaJ, for the rapid differentiation of closely related clinical isolates such as M. avium subspecies avium and Mycobacterium avium subspecies paratuberculosis, and subspecies of M. kansasii.47–51 In 2000, Lee et al. amplified 360-bp fragment within rpoβ gene and digested with restriction enzymes Msp I, Hae III, Sau 3 AI, and Hinc II for differentiation of most of the mycobacteria to the species and subspecies level.49 Besides

Mycobacterium

differentiating among, M. abscessus-M. chelonae-M. fortuitum, M. avium-M. paratuberculosis-M. intracellulare, M. marinum-M. ulcerans, and M. celatum types 1 and 2, rpoβ PCR–RE allowed the differentiation of six subspecies of M. kansasii. PCR-Based Sequencing. PCR-based sequencing has become the gold standard for identification of mycobacterial species. The method consists of PCR amplification of mycobacterial DNA with genus-specific primers and sequencing of the amplicons. The organism is identified by comparison of the nucleotide sequence with reference sequences. Usually, only one sequencing reaction is needed for a definite identification. This method allows for direct detection of mycobacterial species that cannot be grown on conventional laboratory culture media, and several previously unrecognized species have been identified.37,52,53 The target most commonly used is the gene coding for the 16S rRNA. This gene is present in all bacterial species and contains both conserved and variable regions, making it an ideal target for taxonomic purposes. Sequencing of two hyper variable regions of the 16S rRNA gene allows for identification of the majority of mycobacterial species. However, the members of the MTB complex cannot be distinguished. For example, M. kansasii has a sequence identical to that of nonpathogenic species M. gastri.37 Several other target genes viz. The 32-kDa protein,45 the 65-kDa heat-shock protein,54 and the 16S-23S rRNA internal transcribed spacer43 have been well characterized for species identification by sequencing. In 1999, Kim et al. reported that comparative sequence analy­sis of the RNA polymerase gene encompassing the region related to RIF resistance in MTB, is useful for the differentiation of mycobacterial species.52 Loop-Mediated Isothermal Amplification (LAMP). LAMP is a novel, rapid, and simplified NA platform developed by Eiken Chemical Co., Ltd. (Tokyo, Japan).55 Its major advantages are speed, simplicity, and lack of requirement for a thermal cycler, thus facilitating high throughput. The technology uses four different primers specifically designed to recognize six distinct regions on the target gene, and the reaction process proceeds at a constant temperature (isothermal) using strand-displacement reaction. Amplification and detection of the gene products can be completed in a single step, by incubating the mixture of samples, primers, DNA polymerase with strand-displacement activity, and substrates at a constant temperature. Currently, there is limited evidence on the accuracy of LAMP for the detection of TB. Studies are underway to define the test performance and compare its performance against conventional test. Real-Time PCR. Real-time PCR has been successfully applied directly to clinical specimens for rapid detection of mycobacteria and drug-resistance testing. Different probes have been used, like the TaqMan probe, fluorescence resonance energy transfer (FRET) probes, molecular beacons, and bioprobes. The main advantages of real-time PCR techniques are the speed of the test and a lower risk of contamination. Major disadvantages are the requirement

121

for expensive equipment and reagents, as well as the need for skilled technical personnel. Results could be obtained in an average of 1.5–2 h after DNA extraction. Real-time PCR could eventually be implemented in reference laboratories with the required capacity to properly set up the technique and in settings where it can contribute to the management of TB patients. DNA Probe Hybridization. The DNA probe technology for identification of fastidious organisms is still one of the most successful molecular diagnostic procedures worldwide. Accuprobe (Gen-Probe, San Diego, CA), is a commercial method based on species-specific DNA probes that hybridize to rRNA for the identification of limited number of important mycobacteria, including MTB complex, M. avium, M. intracellulare, MAC, M. kansasii, and M. gordonae. The probes have been extensively evaluated in a clinical setting and have shown good sensitivity and specificity with results in 2 h. Overall, DNA probes show >99% sensitivity and specificity for MTB complex isolates and approximately 95% sensitivity and >99% specificity for MAC isolates.56–61 Although rare, some instances of cross-reaction between MTB complex probe and M. terrae and M. celatum isolates and M. avium complex probe with recently described species M. palustre, M. parascrofulaceum, and M. saskatchewanense have been reported.62,63 Line Probe Assays. The line probe assay uses the reverse line blot hybridization technology with differently specific DNA probes immobilized in parallel lines on a paper strip or a nylon membrane. The target DNA is PCR amplified using biotinylated primers and finally incubated with a strip or membrane coated with speciesspecific probes. Finally hybrids are detected through a colorimetric or enhanced chemiluminiscence (ECL) reaction.36,37,64 Three commercial kits for species identification are currently available in the market for detection of mycobacteria and drug-resistance testing: INNO-LiPA, MYCOBACTERIA (Innogenetics NV, Gent, Belgium)65,66; Geno Type Mycobacterium (Hain, Germany)36,37; and Geno Type MTBC (Hain, Germany). Many studies conducted on application of LiPA assay on MTB isolates for detection of RIF resistance showed greater than 95% sensitivity and 100% specificity. Very few studies that applied LiPA directly to clinical specimens had sensitivity ranged from 80%–100%.65,66 The GenoType MDBDR, on the other hand, detects resistance to INH and RIF in culture samples based on the most common mutations in the katG and rpoβ genes, respectively. DNA Arrays. More recently, DNA microarrays or highdensity oligonucleotide arrays (DNA chip) have been applied for species identification of mycobacteria. DNA chip generally comprises a glass surface on which multiple DNA probes with known identities are fixed for molecular hybridization with specific PCR amplified products, which allows the examination of parallel gene expression or genotyping.67 This method allows the simultaneous analysis of thousands of genes in a short assay time and as such is useful for phylogenetic analysis and species identification. For the identification

122

Molecular Detection of Human Bacterial Pathogens

of bacteria, this method may involve the labeling of in vitro RNA transcribed from a target gene from bacteria in specimens, subsequent hybridization of the labeled in vitro transcribed RNA to species-specific oligonucleotide probes on a microarray, and detection of the label, usually by fluorescence. The Affymetrix Genechip, which uses large sets of oligonuleotides that are synthesized rather than spotted onto a glass substrate, has been successfully applied using 16S rRNA genes as a target for the identification of mycobacterium species isolates.68,69 The recently described CombiChip Mycobacteria DrugResistance detection DNA chip is an oligonucleotide microchip coupled with PCR for the detection of resistance to INH and RIF. It was compared with sequencing and drug susceptibility testing in 69 INH and/or RIF resistant and 27 drug-susceptible MTB isolates.70 It allowed identification of 84% of INH resistant isolates based on katG codon 315 and inhA-15 mutations, and 100% of RIF-resistant isolates based on seven codons on rpoβ gene viz. 511, 513, 516, 522, 526, 531, and 533. The overall specificity was 100% and 95.3% for detecting INH and RIF resistance, respectively. Due to the high cost involved, for the time being the use of microarrays for detecting drug resistance in TB is still beyond the reach of most clinical mycobacteriology laboratories. GeneXpert. Xpert(R) MTB/RIF is a test that can simultaneously identify MTB and resistance to rifampicin (RIF). It was developed through a partnership between Cepheid and the Foundation for Innovative New Diagnostics (FIND), the University of Medicine and Dentistry of New Jersey. The system works on the principle of real-time nested PCR and the detection of amplicons by using molecular beacons. It can deliver an accurate diagnosis of the disease in less than 2 h. This system is basically designed for clinicians so that proper treatment can be initiated rapidly.

5

11.2  METHODS 11.2.1  Sample Preparation DNA Extraction. The ability of molecular assays to detect mycobacteria in clinical specimens is dependent on both the target sequence selected and the efficiency of the DNA extraction procedure. The most commonly used methods for DNA extraction are Tris-EDTA (TE) boil extraction, lysis extraction (Infectio Diagnostics, Inc. Quebec, Canada), QIAGEN QIAmp DNA mini kit (QIAGEN, Inc., Valencia, CA), sodium dodecyl sulfate (SDS)-Triton X extraction, and SDS-Triton X plus sonication.

11.2.2  Detection Procedures (i) Standard PCR Principle.  In PCR, DNA is amplified using two primers and one enzyme (Figure 11.1). The repeated thermal cycling of PCR begins with denaturation of the double stranded DNA amplicon followed by primer annealing to target DNA and extension of PCR products by Taq polymerase in presence of excess of deoxynucleoside triphosphates (DNTPs). Each cycle theoretically doubles the number of target nucleic acid molecules; therefore, after 20 cycles, million-fold amplification is achieved, and after 30 cycles, a billion-fold amplification is observed.71 Procedure.  A basic PCR set up requires several components and reagents viz. DNA template, two primers complementary to the 3' ends of each of the sense and antisense strand of the DNA target, Taq polymerase, deoxynucleoside triphosphates (DNTPs), Buffer solution, divalent cations, magnesium or manganese ions, and monovalent cation potassium ions. The PCR is commonly carried out in a reaction volume of 10–200 μL in small reaction tubes (0.2–0.5 mL volumes) in a thermal cycler. The PCR usually consists of a

3

Target Sequence

Step 1: Denaturation 3 5 5

5

3’

Primer 2

3

3 3

Step 2: Annealing

Primer 1

5

5 5 5

3

3 5

Taq DNA Polymerase

Step 3: Extension

5

5

3

5

3

3

5

3

5

Results in Two Copies of Target Sequence

FIGURE 11.1  PCR—Amplification of DNA.

123

Mycobacterium

series of 20–40 cycles. Detection of amplified PCR products can be done by gel electrophoresis. The size of PCR product is determined by comparison with a DNA ladder (molecular weight marker), which contains DNA fragments of known size, run on the gel alongside the PCR products.

design, it enables rapid generation of high quality meaningful data. A suitable pair of primers and probes can be designed based on empirical design guidelines and using primer design softwares. General quantitative real-time PCR protocol is summarized in Figure 11.2.

Note: Incorporation of a negative control, positive control, and internal control is essential with each batch of PCR.

Note: Multiple housekeeping genes should be validated for normalization in each experiment and incorporation of an external standard (alien RNA) is also needed to identify possible PCR inhibitors between treatments and runs.

(ii) Real-Time PCR Principle.  Real-time PCR methods employ fluorescent dyes or fluorophore-containing DNA probes to measure the amount of amplified product. TaqMan probe consists of two types of fluorophores, which are the fluorescent parts of reporter proteins. The probe is attached or unattached to the template DNA, and before the polymerase acts, the quencher fluorophore (usually a long-wavelength colored dye, such as red) reduces the fluorescence from the reporter fluorophore (usually a short-wavelength colored dye, such as green). It does this by the use of fluorescence resonance energy transfer, which is the inhibition of one dye caused by another without emission of a proton. The reporter dye is found on the 5' end of the probe and the quencher at the 3' end. There are two other types of real-time PCR methods, the molecular beacon method and the SYBR® Green method. The molecular beacon method utilizes a reporter probe that is wrapped around into a hairpin. It also has a quencher dye that must be in close contact to the reporter to work. An important difference of the molecular beacon method in comparison to the TaqMan® method is that the probe remains intact throughout the PCR product, and is rebound to the target at every cycle.

(iii) Genotyping Principle.  Genotyping refers to the process of determining the genotype of a MTB complex genome by the use of biological assays. Current methods include PCR DNA sequencing, Allele Specific Oligonucleotide (ASO) probes and hybridization to DNA microarrays or beads. Mycobacterial genotyping technique is used to compare DNA from different sources or to identify a genome by comparison with a known standard. Procedure.  The genotyping laboratories can use three genotyping methods: spoligotyping, Mycobacterium Interspersed Repetitive Units (MIRU) analysis, and IS6110based Restriction Fragment Length Polymorphism (RFLP) analysis, also known as fingerprinting. Spoligotyping and MIRU analysis are based on PCR. Together, these two methods are referred to as the PCR genotyping tests. RFLP. RFLP analysis was performed by the internationally standardized procedure.72 In brief, MTB genomic DNA is extracted and digested with PvuII. The resulting DNA fingerprints are separated by gel electrophoresis, transferred to nylon membrane by the Southern blotting and probed with a horseradish peroxidase-labeled 245 bp sequence of IS6110 DNA probe. Hybridization of probe to a piece of minisatellite

Procedure.  Real-time PCR is an exceptionally powerful research tool. With the correct kit reagent and experimental Primers and Prob Designing

Sequence Specificity BLAST primers to confirm specificity PCR Optimization Annealing Temp Mg2+ conc. Primer conc., Template conc.

Absolute quantification Standard Curve Standards, samples, and negative control in triplicates

Continuous flourescent measurement of PCR product during each cycle of PCR

FIGURE 11.2  General Protocol for Standardization of Quantitative Real-Time PCR.

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Molecular Detection of Human Bacterial Pathogens

DNA results in a dark band on an autoradiograph of the nitrocellulose paper. The resulting patterns are unique to each individual. Spoligotyping. Spoligotyping is generally performed with a commercially available kit (Isogen Bioscience). Briefly, the procedure involves amplification of the direct repeat (DR) region of M. tuberculosis genome by PCR using specific biotinylated primers.73 The amplified products are then hybridized onto a nylon membrane blotted with 43  spacer sequences. The spacer sequences are specific to the DR region and the strains differed in presence or absence of these spacer. The results of hybridization are detected by ECL detection system.72,74 MIRU-VNTR. Mycobacterial interspersed repetitive unit-variable number tandem repeat (MIRU) typing is a MTB complex specific name of a multiple locus variable number of tandem repeats (VNTR) analysis. This method relies on PCR amplification using primers specific for the flanking regions of the VNTRs, and on determination of the sizes of the amplicons, which reflect the numbers of the amplified VNTR copies. Their use provides typing data in a portable numerical format, adequate for studying the global molecular epidemiology of infectious agents.75,76

not yet gained easy access to the routine procedures performed in the clinical mycobacteriology laboratory, especially in low-income TB endemic countries. In these countries with limited resources, the diagnosis of mycobacterial infections is established empirically on clinico-radiological basis or only by sputum AFB smear examination. Due to lack of facilities most laboratories in disease endemic countries do not identify the mycobacteria by culture and infections caused by NTM are underdiagnosed. Today, with techniques and equipment becoming increasingly complex and expensive, it is difficult to upgrade local laboratories to perform these assays. As specimen delivery and communication of results can be rapidly and easily achieved, utilization of reference laboratories for rarely performed sophisticated molecular tests would be a more practical approach.

Note: Genotyping of MTB isolates is useful in TB control for confirming the dynamics of TB outbreaks, laboratory cross-contamination, and identified MTB strains, transmission sites, and accurate epidemiologic links.



11.3  C  ONCLUSION AND FUTURE PERSPECTIVES It is important to realize that there is no “stand alone” assay for the identification of mycobacteria especially NTM. Many new species may not be recognized in all assays. This plethora of species poses an additional challenge for the clinical mycobacteriology laboratory to provide timely services. Furthermore, some of the species (e.g., M. haemophilum, M.  genavense) require special growth conditions, necessitating an exquisite collaboration between the clinician requesting the test and the laboratory professional performing the test. To overcome these shortcomings of conventional methods, in recent years molecular techniques have become increa­ singly popular, as they are rapid, reliable, highly sensitive, and specific and can be used for large number of samples. Accuprobe (Gen-Probe, San Diego, Calif.) and PCR-based assays are more widely used in developed countries for species identification of mycobacteria than phenotypic tests and have significantly improved turnaround time and commonly used. However, results of these tests should be correlated with the clinical presentation and the clinician should notify the laboratory when results are not consistent. The DNA microarray method holds promise for the future because it is easy to perform, can be readily automated, and allows for identification of a large number of mycobacterial species in one reaction. However, due to the requirement of additional equipment and trained personnel, these methods are expensive and have

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Mycobacterium 14. Katoch, V.N., Infections due to non tuberculous mycobacteria (NTM), Indian J. Med. Res., 120, 290, 2004. 15. Graybill, J.R., et al., Disseminated mycobacteriosis due to Mycobacterium abscessus in two recipients of renal homograft, Am. Rev. Respir. Dis., 109, 4, 1974. 16. Iseman, M.D. et al., Disease due to Mycobacterium aviumintracellulare, Chest, 87, 139S, 1985. 17. ATS guidelines 2007, David E. Griffith, Timothy Aksamit, Barbara A. Brown-Elliott. An Official ATS/IDSA statement: Diagnosis, treatment, and prevention of nontuberculous mycobacterial diseases, Am. J. Respir. Crit. Care Med., 175, 367, 2007. 18. Collins, C.H. et al., Mycobacterium marinum infectious in man, J. Hyg. (Lond)., 94, 135, 1985. 19. Grange, J.M., Infections caused by opportunist mycobacteria: A review, J. Royal Soc. Med., 79, 226, 1986. 20. Jackson, P.G. et al., Injection abscesses due to Mycobacterium chelonae occurring in a diabetic patient, Tubercle, 62, 277, 1981. 21. Borghans, J.G., and Standford, J.L., Mycobacterium chelonae in abscesses after injection of diphtheria-pertussis-tetanuspolio vaccine, Am. Rev. Respir. Dis., 107, 1, 1973. 22. Kuritaky, J.N. et al., Sternal wound infections and endocarditis due to organisms of the Mycobacterium fortuitum complex, Ann. Intern. Med., 98, 938, 1983. 23. Gangadharam, P.R.J., Lanier, J.D., and Jones, D.E., Keratitis due to Mycobacterium kansasii, Tubercle., 59, 55, 1978. 24. Moulsdale, M.T. et al., Infection by Mycobacterium haemophilum, a metabolically fastidious acid-fast bacillus, Tubercle., 64, 29, 1983. 25. Cohen, R.J. et al., Occult infections with M. intracellulare in bone-marrow biopsy specimens from patients with AIDS, N. Engl. J. Med., 308, 1475, 1983. 26. Graybill, J.R. et al., Disseminated mycobacteriosis due to Mycobacterium abscessus in two recipients of renal homograft, Am. Rev. Respir. Dis., 109, 4, 1974. 27. Gordin, F., and Slutlin, G., The validity of acid-fast smears in the diagnosis of tuberculosis, Arch. Pathol. Lab. Med., 114, 1025, 1990. 28. Kent, and Kubica, G.P., Public health mycobacteriology a guide for level III lab. Atlanta, GA: U.S. Department of health and human services, Public health services. Center for disease control, 64, 1985. 29. Mole, R.J., and WO’C Maskell, T., Review phage as a diagnostic—the use of phage in TB diagnosis, J. Chem. Technol. Biotechnol., 76, 683, 2001. 30. Shenai, S., Rodrigues, C., and Mehta, A.P., Evaluation of a new phage amplification technology for the rapid diagnosis of tuberculosis, Int. J. Med. Microbiol., 20, 194, 2002. 31. Kalantri, S. et al., Bacteriophage-based tests for the detection of Mycobacterium tuberculosis in clinical specimens: A systematic review and meta-analysis, BMC Infect. Dis., 5, 59, 2005. 32. Jacob, W.R. et al., Rapid assessment of drug susceptibilities of Mycobacterium tuberculosis by means of luciferase reporter phages, Science, 260, 819, 1993. 33. Butler, W.R., and Kilburn, J.O., Identification of major slowly growing pathogenic Mycobacteria and Mycobacteria gordonae by high performance liquid chromatography of their mycolic acids, J. Clin. Microbiol., 26, 50, 1988. 34. Butler, W.R., and Guthertz, L.S., Mycolic acid analysis by high-performance liquid chromatography for identification of Mycobacterim species, Clin. Microbiol. Rev., 14, 704, 2001.

125 35. Eing, B.R., Becker, A., Sohns, A., and Ringelmann, R. Comparison of Roche Cobas Amplicor Mycobacterium tuberculosis assay with in-house PCR and culture for detection of M. tuberculosis, J. Clin. Microbiol., 36, 2023, 1998. 36. Pai, M., Kalantri, S., and Dheda, K., New tools and emerging technologies for the diagnosis of tuberculosis: Part II. Active tuberculosis and drug resistance, Expert Rev. Mol. Diagn., 6, 423, 2006. 37. Tortoli, E., and Palomino, J.C., New diagnostic methods. In Tuberculosis, Chapter 14, p. 441, 2007. 38. Bergmann, J.S., and Woods, G.L., Clinical evaluation of the BD-ProbeTec Strand Displacement Amplification Assay for rapid diagnosis of tuberculosis, J. Clin. Microbiol., 36, 2766, 1998. 39. Piersimoni, C., and Scarparo, C., Relevance of commercial amplification methods for direct detection of Mycobacterium tuberculosis complex in clinical samples, J. Clin. Microbiol., 41, 5355, 2003. 40. Chengtao, H., and Weishilbourn, R.M., The polymerase chain reaction (PCR) is the most widely used technique for the study of DNA. Applications of PCR have been extended significantly by the development of long PCR, Curr. Issues Mol. Biol., 1, 77, 1999. 41. Johansen, I.S., Rapid diagnosis of mycobacterial diseases and their implication on clinical management, Dan. Med. Bull., 53, 28, 2006. 42. Flores, L.L. et al., In-house nucleic acid amplification tests for detection of Mycobacterium tuberculosis in sputum specimens meta analysis and meta-regression, BMC Microbiol., 5, 55, 2005. 43. Roth, A. et al., Differentiation of phylogenetically related slowly growing mycobacteria based on 16S-23S rRNA gene internal transcribed spacer sequences, J. Clin. Microbiol., 36, 139, 1998. 44. Verma, A., Rattan, A., and Tyagi, J.S., Development of 23S rRNA based PCR for the detection of mycobacteria, Indian J. Biochem. Biophys., 31, 288, 1994. 45. Soini, H., Bottger, E.C., and Viljanen, M.K., Identification of mycobacteria by PCR-based sequence determination of the 32-kilodalton protein gene, J. Clin. Microbiol., 32, 2944, 1994. 46. Cave, M.D. et al., Stability of DNA fingerprint pattern produced with IS6110 in strains of Mycobacterium tuberculosis, J. Clin. Microbiol., 32, 262, 1994. 47. Telenti, A. et al., Rapid identification of mycobacteria to the species level by polymerase chain reaction and restriction enzyme analysis, J. Clin. Microbiol., 31, 175, 1993. 48. Vaneechoutte, M. et al., Identification of Mycobacterium species by using amplified ribosomal DNA restriction analysis, J. Clin. Microbiol., 31, 2061, 1993. 49. Lee, H. et al., Species identification of mycobacteria by PCR– restriction fragment length polymorphism of the rpoβ gene, J. Clin. Microbiol., 38, 2966, 2000. 50. Roth, A. et al., Novel diagnostic algorithm for identification of mycobacteria using genus-specific amplification of the 16S-23S rRNA gene spacer and restriction endonucleases, J. Clin. Microbiol., 38, 1094, 2000. 51. Magalhaes, V.D. et al., Reliability of hsp65-RFLP análisis for identification of species in cultured strains and clinical specimens, J. Microbiol. Methods, 49, 295, 2002. 52. Kim, B.J. et al., Identification of mycobacterial species by comparative sequence analysis of the RNA polymerase gene (rpoB), J. Clin. Microbiol., 37, 1714, 1999.

126 53. Kapur, V. et al., Rapid Mycobacterium species assignment and unambiguous identification of mutations associated with antimicrobial resistance in Mycobacterium tuberculosis by automated DNA sequencing, Arch. Pathol. Lab. Med., 119, 131, 1995. 54. McNabb, A. et al., Assessment of partial sequencing of the 65-kiodalton heat shock protein gene (hsp 65) for routine identification of mycobacterium species isolated from clinical sources, J. Clin. Microbiol., 42, 3000, 2004. 55. Zhu R.Y., et al. Use of visual loop-mediated isothermal amplification of rimM sequence for rapid detection of Mycobacterium tuberculosis and Mycobacterium bovis, J Microbiol. Methods., 78(3), 339, 2009. 56. Bull, T.J., and Shanson, D.C. Evaluation of a commercial chemiluminescent Gen Probe system “AccuProbe” for the rapid identification of mycobacteria including “MAIC X”, isolated from blood and other sites, from patients with AIDS. J. Hosp. Infect., 21, 143, 1992. 57. Tortoli, E. et al. Evaluation of a commercial DNA probe assay for the identification of Mycobacterium kansasii, Eur. J. Clin. Microbiol. Infect. Dis., 13, 264, 1994. 58. Tortoli, E., Simonetti, M.T., and Lavinia, F., Evaluation of reformulated chemiluminescent DNA probe (Accuprobe) for culture identification of Mycobacterium kansasii, J. Clin. Microbiol., 34, 2838, 1996. 59. Saito, H. et al., Identification and partial characterization of Mycobacterium avium and Mycobacterium intracellulare by using DNA probes, J. Clin. Microbiol., 27, 994, 1989. 60. Gonzales, R., and Hanna, B.A., Evaluation of Gen-Probe DNA hybrodozatopn system for the identification of Mycobacterium tuberculosis and Mycobacterium avium-intracellulare, Diagn. Microbiol. Infect. Dis., 8, 69, 1987. 61. Shenai, S., Rodrigues, C., and Mehta, A., Rapid diagnosis of tuberculosis using transcription mediated amplification, Indian J. Med. Microbiol., October, 19, 184, 2001. 62. Torkko, P. et al., Mycobacterium palustre sp nov., a potentially pathogenic slow-growing mycobacterium isolated from veterinary and clinical specimens, and Finnish stream water, Int. J. Syst. Evol. Microbiol., 52, 1519, 2002. 63. Turenne, C.Y. et al., Mycobacterim, parascrofulaceum sp, nov., a novel slowly growing, scotochromogenic clinical isolates related to Mycobacterium interjectum and Accuprobe positive for Mycobacterium avium complex, Int. J. Syst. Evol. Microbiol., 54, 659, 2004.

Molecular Detection of Human Bacterial Pathogens 64. Shenai, S., Rodrigues, C., and Mehta, A., Rapid speciation of 15 clinically relevant mycobacteria with simultaneous detection of resistance to rifampicin, isoniazid and streptomycin in M. tuberculosis complex, Int. J. Infect. Dis., 13, 46, 2009. 65. Tortoli, E., Mariottini, A., and Mazzarelli, G., Evaluation of INNO-LiPA MYCOBACTERIA V2: Improved reverse hybridization multiple DNA probe assay for mycobactrial identification, J. Clin. Microbiol., 41, 4418, 2003. 66. Martin, A., and Porteals, F., Drug resistance and drug resistance detection. In Tuberculosis, Chapter 19, p. 635, 2007. 67. Fukushima, M. et al., Development of DNA microarray method to detect and identify pathogenic bacteria, Jpn. J. Electrophoresis, 45, 235, 2004. 68. Troesch, A. et al., Mycobacterium species identification and rifampicin resistance testing with high-density DNA probe arrays, J. Clin. Microbiol., 37, 1079, 1999. 69. Fukushima, M. et al., Detection and identification of Mycobacterium species isolates by DNA microarray, J. Clin. Microbiol., 41, 2605, 2003. 70. Caoli, J.C. et al., Evaluation of the TB-Biochip oligonucleotide microarray system for rapid detection of rifampin resistance in Mycobacterium tuberculosis, J. Clin. Microbiol., 44, 2378, 2006. 71. Erlich, H.A., PCR Technology: Principles and Applications for DNA Amplifications, Stockton Press, New York, 1989. 72. van Embden, J.D. et al., Strain identification of Mycobacterium tuberculosis by DNA fingerprinting: Recommendations for a standardized methodology, J. Clin. Microbiol., 31, 406, 1993. 73. Kamerbeek, J. et al., Simultaneous detection and strain differentiation of Mycobacterium tuberculosis for diagnosis and epidemiology, J. Clin. Microbiol., 35, 907, 1997. 74. Groenen, P.M.A. et al., Nature of DNA polymorphism in the direct repeat cluster of Mycobacterium tuberculosis: Application for strain differentiation by a novel typing method, Mol. Microbiol., 10, 1057, 1995. 75. Supply, P. et al., Automated high-throughput genotyping for study of global epidemiology of Mycobacterium tuberculosis based on mycobacterial interspersed repetitive units, J. Clin. Microbiol., 39, 3563, 2001. doi: 10.1128/ JCM.39.10.3563–3571.2001. 76. Klevytska, A. et al., Identification and characterization of variable-number tandem repeats in the Yersinia pestis genome, J. Clin. Microbiol., 39, 3179, 2001.

12 Nocardia Veronica Rodriguez-Nava, Frédéric Laurent, and Patrick Boiron CONTENTS 12.1 Introduction...................................................................................................................................................................... 127 12.1.1 Classification......................................................................................................................................................... 127 12.1.2 Epidemiology........................................................................................................................................................ 128 12.1.3 Clinical Features and Pathogenesis...................................................................................................................... 129 12.1.3.1 Localized Pulmonary Nocardiosis........................................................................................................ 129 12.1.3.2 Extrapulmonary Localized Nocardiosis................................................................................................ 129 12.1.4 Diagnosis.............................................................................................................................................................. 130 12.1.4.1 Phenotypic Techniques.......................................................................................................................... 130 12.1.4.2 Molecular Techniques.............................................................................................................................131 12.2 Methods............................................................................................................................................................................ 134 12.2.1 Sample Preparation............................................................................................................................................... 134 12.2.2 Detection Procedures............................................................................................................................................ 134 12.2.2.1 Genus-Specific Identification................................................................................................................. 134 12.2.2.2 Species-Specific Identification............................................................................................................... 134 12.3 Conclusions and Future Work........................................................................................................................................... 134 References.................................................................................................................................................................................. 135

12.1  INTRODUCTION Bacteria of the Nocardia genus are ubiquitous in the environment. They have been isolated in soil, plants, and compost, as well as in humans. The infections they cause generally originate from the inhalation of the bacteria or by their contamination of wounds. Their proliferation within the body gives rise to a localized or disseminated granulomatous and suppurative infection, referred to as nocardiosis.1 Nocardia infections have long been considered to be rare, especially since the diagnosis of nocardiosis, whether direct, indirect, or immunologic, has long been out of the reach of a large number of laboratories. In recent years, the situation has dramatically changed. The number of patients suffering from nocardiosis worldwide is constantly increasing. Although a majority of patients have risk factors (generally an immunosuppressive situation), it must be noted that 30% of cases were observed in subjects who were apparently immunocompetent. Due to the nature and seriousness of the infection, the accurate and rapid detection and identification of such bacteria, as well as an assessment of their sensitivity levels to antibiotics, are essential. Diagnostic methods to replace conventional ones were proposed to microbiology laboratories in order to help them more quickly detect and identify Nocardia species whose number has tripled within the past few years

and which display different epidemiological and pathogenic characteristics.

12.1.1  Classification Bacteria with positive Gram staining form one of the ten major suprageneric groups2 which is further divided into two principle groups depending on DNA base composition. The actinomycete group consists of bacteria whose DNA is rich in guanine (G) and cytosine (V), characterized by a G + C% > 55. The use of 16S rRNA sequencing techniques has allowed for the identification of phylogenetic relations between prokaryotes groups.3 The standard chemical identification process of actinomycetes relies on Lechevalier’s chemotaxonomic system,4 which allows one to distinguish between 10 chemotypes. This system is based on the presence of parietal amino acids [2,4-diaminobutyric acid, meso-2,6-diaminopimelic acid (meso-DAP), glycine, lysine, ornithine, aspartic acid] and parietal lipids (fatty acids, phospholipids, menaquinones, mycolic acids), as well as cellular sugars (arabinose, galactose, madurose, xylose) whose single or combined predominant presence provides a possible indication of a particular genus. Within the past two decades, improvements in methods for the identification of bacteria, such as chemical analysis 127

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techniques (major cellular and membranous component) and molecular biology techniques (G + C%, DNA–DNA homologies, DNA–RNA, 16S rRNA sequencing), have lead to fundamental changes in the classification and identification of such bacteria. The Nocardia genus shares two types of features with other actinomycete genera. The first is morphological: its fragile transient primary (aerial) mycelium is split into bacillary and coccoid elements, which makes Nocardia bacteria resemble other nocardioform actinomycete genera, according to Lechevalier definition (Rhodococcus, Faenia, Pseudonocardia, Saccharomonospora, Saccharopoly-spora, Actinopolyspora, Amycolata, Amycolatopsis, Oerskovia, Promicromonospora, Nocardioides, Intrasporangium). Aside from Rhodococcus, these genera resemble the Nocardia genus neither from a phylogenetic nor biochemical point of view.4 In terms of phylogenetic and biochemical features, the Nocardia genus resembles the Rhodococcus, Gordonia, Tsukamurella, Dietzia, and Skermania genera forming the Nocardiaceae family and the Mycobacterium and Corynebacterium genera within a suprageneric group of nocardioform actinomycetes, according to the definition of Goodfellow.5,6

12.1.2  Epidemiology Habitat. Nocardia bacteria live as saprophytes in the soil and can be found in decomposing vegetable matter, in fresh or salt water, and in atmospheric dust. They are vastly present in soil, where as a saprophyte they contribute to the decomposition of vegetal, organic matter. These bacteria can be easily isolated in fresh and salt water, compost, plant matter, or atmospheric dust. Several authors have demonstrated the abundance of actinobacteria in caves,7–9 which leads to the conclusion that such a habitat is particularly prosperous for the development of this group of bacteria. Diverse new species of the Nocardia genus described in the recent years, among which N. jejuensis, N. speluncae, and N. altamirensis were isolated from subterraneous environments. The identification of these species indicates that such environments constitute an ecological niche with a high potential for the study of biodiversity and the description of new species. Nocardia bacteria are not part of the standard microbial flora of humans. Although the presence of Nocardia on the skin and inside the respiratory tract has been reported, their isolation usually indicates a patent or latent infection, particularly among immunosuppressed patients, and must therefore undergo clinical analysis.10,11 Contamination Modes. The contamination by Nocardia species generally occurs through the inhalation of spores, originating from telluric bacteria or occasionally from a skin trauma. A metastatic presence in the central nervous system is in most cases the consequence of a hematogenous dissemination from an initial pulmonary or cutaneous contamination.10

Molecular Detection of Human Bacterial Pathogens

Entry through the digestive tract is much more rare and leads to isolated abdominal forms. Interhuman transmission between immunosuppressed patients has been reported. Several species of the Nocardia genus were found in domestic and hospital air and dust. This observation may be an indication of a risk of nosocomial infections although only a few cases have been reported.12 Influence of Age and Gender. Nocardia infections mostly occur in adults aged 60–80 years and with no significant difference between the genders. Prevalence and Incidence of the Disease. Five hundred to one thousand cases of nocardiosis were diagnosed annually in the in the United States during the 1970s,13 but the actual incidence was probably higher due to a low “suspicion index” (inaccurate diagnosis techniques, confusion with other infections, and intercurrent pathologies sometimes masking diagnosis). A more recent epidemiological study of nocardiosis conducted by Saubolle et al. (2003)14 in the state of Arizona reported 455 cases diagnosed between 1998 and 2002, the most frequent etiologic agent being the N. asteroides, present in 307 cases. A survey conducted in France between 1987 and 1990 reported 150–250 new cases annually.15 Nearly 1000 strains were sent to the French Nocardiosis Observatory between 2000 and 2007. More than 75% (880 cases) belong to the actinobacteria family. Among these, 607 cases of nocardiosis originating from all regions of France (mainland and overseas) were identified. The most frequent species found in 75% of the clinical isolates are N. farcinica (26%), N. nova (20%), N. abscessus (18%), and N. cyriacigeorgica (12%).16 The apparent increase in incidences of nocardiosis can be explained by the aging of the general population and by the use of aggressive immunosuppressive therapeutics. A better knowledge of nocardiosis and improvements in diagnostic methods are also possible explanations for the increases of diagnosed cases. Predisposition Factors. Patients presenting primary or secondary immunosuppression are particularly predisposed to Nocardia infections. Long-term corticotherapy appears to be the main factor of predisposition as it is found in nearly a third of all patients. Transplantations and hematology also constitute major risk factors. Neoplasia, and particularly those affecting the tracheobronchial tree (oto-rhino-laryngological and pulmonary cancers), bronchial dilation, and HIV also constitute predisposing factors. No predisposing factor was found in one-third of all cases.10 A number of debilitating factors were described: tuberculosis, silicosis, disseminated acute erythematous lupus, periarteritis nodosa, polyarthralgia rheumatica, pemphigoid, asthma, emphysema, bronchial and obstructive chronic bronchopneumopathy, pulmonary alveolar proteinosis, sarcoidosis, diabetes, malnutrition, intestinal lipodystrophy, colite ulcerosa, chronic alcoholism, toxicomania, and pregnancy.17

Nocardia

12.1.3  Clinical Features and Pathogenesis 12.1.3.1  Localized Pulmonary Nocardiosis Clinical Presentation. The lung is the most commonly affected organ, touched in 60%–80% of the cases.18 Pulmonary nocardiosis generally takes the form of chronic pneumopathy but can become acute following a rapid degra­ dation. A pulmonary lesion is a localized pneumopathy surrounded by an inflammatory response. Occasionally, an abscess can form and progressively spread. The clinical expression of nocardiosis may also include forms such as bronchitis or pharyngitis. The clinical pulmonary signs are polymorphous. Some evoke a bronchial syndrome, with dyspnea, thoracic pain of pleuropulmonary origin, and possible respiratory distress. Other symptoms can be observed, including fever, shivering, asthenia, anemia, anorexia, general state alteration, and night sweats. Hemoptysis is rare and expectorating pulmonary purulence or mucupurulence (often of the color green) can appear. Nocardiosis can provoke a mediastinitis with a large venous obstruction and a superior caval syndrome. Biology. Biological signs are neither obvious nor specific. A polynuclear neutrophils hyperleukocytosis (whose importance is unrelated to the seriousness of the infection) can be found occasionally. However, a moderate anemia associated to other inflammatory signs is generally observed. Radiology. Radiological signs of pulmonary nocardiosis, highly polymorphous and unspecific, do not allow for a diagnosis. Nocardiosis can lead to a wide variety of radiological images with one or both lungs being affected, most commonly with apical localization: infiltrates, segmented, lobar or diffuse condensation, necrotizing pneumopathy, small or large abscesses, pleural effusion, cavities of all sizes, single or multiple nodular images with military aspect, fistulous tracts (in case of the spread of the infection to the thoracic wall), and, more rarely, hilary lesions, adenopathies, or calcifications. Differential Diagnosis. The clinical differential diagnosis has to be conducted for all typical or atypical pneumopathies, whether of bacterial, mycobacterial, viral, mycotic, or toxic origin, whether particular or medicinal. The differential diagnosis must exclude the pulmonary expression of a systematic illness, as well as primary or secondary neoplastic disorders. These diagnostic difficulties may in some cases lead to a pulmonary biopsy that can help distinguish between various hypotheses. Evolution. The Nocardia infection has a tendency to spread from its primary pulmonary site to other tissues through the lymphatic or circulatory system. The central nervous system constitutes the most frequent metastatic localization. The skin and subcutaneous tissues are less frequently touched, along with the pleural cavity and the thoracic wall, the eye, the liver, the lymphatic ganglions and other sites: intestines, bones, heart, pericardium, endocardium, kidneys, endocrine glands, and so forth.

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The pulmonary infection is sometimes asymptomatic and nocardiosis is revealed through pulmonary radiography or through the emergence of one or several infectious, metastatic sources. 12.1.3.2  Extrapulmonary Localized Nocardiosis (i) Neuromeningeal Forms  They are most frequently secondary with a pulmonary source.19 Clinical Presentation. The infection of the central nervous system generally takes the form of one or several abscesses that can be situated anywhere in the brain (cerebellum, pons, rachidian bulb, Gasser’s ganglion) with a tendency to form satellite extensions. A meningitic infection, whether acute or chronic, generally originates from a rupture of a cerebral abscess in the ventricular system. Pure meningitis without the presence of any cerebral abscesses is more rarely observed. The clinical situation results from the intracerebral spread of the infection, with variable signs and focal symptoms depending of the affected region of the brain: cephalea (often chronic, preceding the emergence of other signs by several weeks or months, and being the most frequent symptom), nausea and vomiting (resulting from an intracranial hypertension), neck stiffness, photophobia (more specific to an meningitic infection), sensitive or motor (most commonly hemiparesis), possibly associated with aphasia or dysarthria, sensory disorder: visual (dilopia, hemianopsia) or auditory, affected cranial nerves, convulsive crisis (signs of cortical irritation), vertigo, instability, ataxia (in case of cerebellar affection), alteration of the superior functions (ranging from confusion and amnesia to coma), behavioral disorder: irritability and frontal syndrome. It must be emphasized that in a number of cases of nocardiosis in cerebral sites, no patent neurological symptoms are found and only a systematic search through imaging will allow one to identify this type of localization. When such a systematic search was conducted, a cerebral localization was identified in more than a third of all patients. Biology. A normal cerebrospinal fluid (CSF) is common in Nocardia infections. However, in cases of meningeous infection, the examination of the CSF may be indicative of an increase in white blood cells (mostly polynuclear neutrophils), a hypoglycorrhachia, and a hyperproteinorachia. Imaging. Cerebral tomodensitometry allows for the diagnosis of the intracerebral spread of the infection as to localization, size, and number. In cerebral nocardiosis, it is essential to carry out imaging after the intravascular injection of a contrast-enhancing agent. In 80% of all cases, cerebral abscesses are seen in the form of concentric circles with a hypodense central zone and peripheral halo. Magnetic resonance imaging (MRI) allow for the identification of the liquid nature of the lesion. Differential Diagnosis. The images obtained (abscess displaying a smooth capsule of uniform thickness and hypodense content) may be compatible with other lesions, mainly a glioma (in which case the central zone appears to be denser), a primitive or metastatic tumor (in which case the

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edge is more regular), a cerebrovascular accident, or multiple sclerosis. Therefore, MRI imaging may turn out to be insufficient for differentiating an abscess from another liquid tumor. In this case, Indium-111 leucocyte scintigraphy may be very helpful. When one or several cerebral abscesses are associated with an extra cerebral source of infection (in particular pulmonary), several other etiologic agents may be suspected: Pneumococcus, Streptococcus, Bacteroides, Mycobacterium tuberculosis, Cryptococcus, Coccidioides, Histoplasma, Blastomyces, Aspergillus. The sterility of the CRF is an argument in favor of nocardiosis. (ii) Cutaneous and Subcutaneous Infections  Cutaneous nocardiosis can be observed in two different epidemiologic and clinical situations. Primitive, cutaneous nocardiosis occurs following a traumatism in health patients. It generally lies on exposed parts of the skin. The lesions are acute or subacute and may resemble pyodermatitis, cellulite, subcutaneous abscess, or lymphangitis. There is no systemic dissemination of the infection and a spontaneous healing of the lesions is possible. Secondary cutaneous nocardiosis is observed primarily in immunosuppressed patients and manifests itself though single or multiple subcutaneous abscesses that can fistulate to the skin, or through cellulites. It originates (in 10%–15% of cases) from hematogenous spread of a source, and is often pulmonary. It may also originate from a direct extension of a pulmonary source through the thoracic wall. (iii) Other Sites of Infection  Almost all organs can be touched secondarily, each giving rise to particular clinical signs: the eye (keratitis, dacryocystitis, chorioretinitis, endophthalmitis), liver (abscess), pancreas (abscess, even pancreatitis), heart (pericarditis, endocarditis), bones (osteitis, osteomyelitis, arthritis), kidneys, suprarenal glands, spleen, intestine, peritoneum (particularly among dialyzed patients), lymphatic ganglions, thyroid, ear canal, amygdales, pharynx, buccal cavity, and trachea (particularly through contamination following transtracheal aspiration).

12.1.4  Diagnosis A diagnosis of nocardiosis must be evoked (along with common bacterial and viral infections, tuberculosis, and fungal opportunistic infection) in cases of febrile, suppurative disease or in the presence of a pathological pulmonary radiographic image for patients whose immune system was altered by certain therapeutic agents or by an underlying pathology.20,21 The diagnosis of nocardiosis relies on bacteriologic identification in patient samples occasionally obtained using intrusive methods. The histology may also be informative.

Molecular Detection of Human Bacterial Pathogens

12.1.4.1  Phenotypic Techniques The identification of Nocardia and the diagnosis of nocardiosis represent great difficulties for nonspecialized microbiological laboratories.22,23 About 30 years ago, conventional identification techniques were developed: strict aerobic respiratory type; macroscopic and microscopic morphology [presence of ramified, gram-positive filaments that split into bacillary and coccoid elements and produce (rarely) short chain conidia]; and chemotaxonomic, biochemical, and physiological characteristics.24 Microscopy. Thin smears prepared from the pure pathologic mixture or from centrifugation sediment as well as the juxtaposition of biopsies can be performed for a microscopic examination. The presence of ramified filaments can be observed, although in a majority of cases it is difficult to detect the often rare bacteria through direct observation. Nocardia display diverse morphological characteristics. The classical appearance is the ramified filament form, but Nocardia filaments are fragile and often break into shorter fragments during the sampling or smearing process. The “zigzagging” aspect of the ramified filaments can be observed in a low-brightness phase-contrast direct examination. The Gram staining is positive, but often weakly and irregularly so (dotted, stained, or striped appearance). Upon Ziehl–Neelsen staining (discoloration by 1% or 2% sulfuric acid during 1 min), the acido-alcohol resistance may vary. Such a partially acid-alcohol fast character allows one to differentiate between Nocardia and bacteria of the Actinomyces and Streptomyces genera. Gomori staining (methenamine silver) may be used to look for Nocardia during histopathological examination, but also stains fungus and Pneumocystis jiroveci. These sole morphological features in an aerobic microorganism have long been considered sufficient to establish the identification of Nocardia. To date, these criteria are not considered sufficient to exclude other bacteria, and more reliable criteria (i.e., analysis of the cellular and parietal constituents) are now required. A negative microscopic examination does not rule out a nocardiosis diagnosis and may be followed by a positive culture if the incubation is sufficiently long.22 Culture. Culturing is easy on substrates free of antibiotics (5% sheep blood agar, enriched blood agar, BCP medium, BACTEC® or BACTalertEC® systems) as well as culture substrates for fungus or mycobacteria (Sabouraud, LöwensteinJensen, Bennett). The use of a selective paraffin medium or a normal or selective BCYE (buffered charcoal yeast extract) medium may eliminate the conventional fast-growing bacteria, which occasionally mask the presence of Nocardia. Cultures are incubated at a temperature ranging from 32°C to 37°C in aerobiosis or possibly in the presence of 5%–10% of CO2. The growth of colonies generally takes between 4 and 5 days but may take up to 2 weeks. Routinely, the colonies are flat or slightly dome shaped, often with a corrugated surface, cerebriform, and covered within a few days with a short duvet of aerial mycelium

Nocardia

that masks the color of the colony and confers it a chalky white appearance. The colonies are encrusted in the agar gel (solid medium). Certain colonies appear to be pigmented, their color ranging from yellowish (or even whitish) beige to orange or red. Their consistency is generally firm although crumbly. Upon microscopic examination Nocardia appear to be ramified and split into bacillary or even coccoid elements within 24–48 h. Biochemical Characterization. Several biochemical properties of Nocardia are useful for their characterization. These include the substrate decomposition profiles (tyrosine, xanthine, uric acid, adenine, hypoxanthine, casein, urea, and testosterone), the growth capacity on carbon-rich substrates (citrate, acetamide, rhamnose, mannitol, sorbitol, etc.), enzymatic activity (arylsulfatase, catalase, acetamidase, galactosidase, urease, etc.), and sensitivity characteristics or resistance to antibiotics (ampicillin, carbenicilline, amoxicillin + clavulanic acid, cefamandole, kanamicine, etc.). The analysis of chemotaxonomical markers such as fatty acids, mycolic acid, or menaquinones is based on the system of Lechevalier et al.25,26 and Goodfellow et al.,6 and essentially allows the identification of a new type of Nocardia. Being laborious and expensive to carry out, this is limited to specialized laboratories. Phenotypical techniques based on the macroscopic and microscopic features of colonies as well as biochemical characteristics allow one to identify the Nocardia species most responsible for infection but fail to bring new species to light. In addition, given the great variety of phenotypical tests applied to Nocardia and described in the literature, no precise choice has been generally agreed upon. Today, they only complement molecular diagnostic techniques for Nocardia, or are limited to a premature diagnosis, according to the minimal criteria for orientation.24 12.1.4.2  Molecular Techniques The emergence of molecular identification methods has enabled the development of several techniques for definitive identification of members of the Nocardia genus.27–30 DNA Hybridization. DNA hybridization is one of the molecular bacteriological methods useful for the identification of microorganisms. Brownell et al.31 developed a system of probes for the identification of N. asteroides from a genomic bank obtained with a cloned GUH-2 strain in the Lambda EMBL3 vector. This work has not benefited from real development, and no other technique of this type has since been proposed. Ribotyping. The ribotyping technique was evaluated by Laurent et al.32 for the Nocardia genus. The comparative study of restriction profiles of 21 strains belonging to complex N. asteroides confirmed both the heterogeneity of this species and the high level of discrimination and reproducibility of the technique for the identification of isolates belonging to this complex. Nevertheless and despite its promising character, ribotyping has never seen diagnostic application due to its long and fastidious implementation.

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In 2007, Kalpoe et al.33 developed a typing technique based on molecular epidemiology (Amplified Fragment Length Polymorphism, AFLP) in order to study a possible epidemic, observed in the renal transplant service of the Leiden Medical Center in the Netherlands, caused by the N.  farcinica species. They compared this technique with others, such as RAPD (Randomly Amplified Polymorphic DNA) and pulsed field electrophoresis. Results showed that the AFLP analysis seems to be an effective alternative to traditional and poorly reproducible methods, such as RAPD for epidemiological typing of N. farcinica strains. PCR–Restriction Fragment Length Polymorphism (PCR–RFLP). The technique for the identification of actinomycetes of medical interest used by Steingrube et al.29,34 based on the number of restriction sites, their positions and the sizes of fragments generated from amplified product of a short segment (440 base pairs) of the gene coding for the heat shock protein (hsp65) and digested by the enzymes (Bst EII, Msp I, Hinf I, and Bsa HI) leading to the establishment of a decision tree for identification was reevaluated and applied to the new taxonomical context of Nocardia (43 species tested). The results clearly showed its inability to distinguish the ensemble of Nocardia species known to date. This technique can nevertheless distinguish without ambiguity species of clinical interest if it is complemented by appropriate phenotypical tests. This method is, therefore, useful by default.35 The work of Laurent et al.27 has enabled the selection of primers within the 3′ region of the ribosomal 16S DNA of Nocardia that allow the specific amplification of a fragment of 596 base pairs for strains belonging to the Nocardia genus. The specificity of the amplified fragment of 596 base pairs can be verified thanks to the position of restriction sites of two enzymes specific to the Nocardia genus: all Nocardia strains tested showed a restriction site specific to the Mnl I enzyme and no restriction site for the enzyme, SacI. This technique is directly applicable to clinical samples (bronchoalveolar lavage, deep puss, biopsies, CSF or puncture of cerebral abscess) without the need for cell cultures.28 With the same aim as Steingrube et al., Conville et al.,30 have proposed a PCR–RFLP technique based on the part of the gene coding the 16S rRNA for the identification of the species of the Nocardia genus. Their goal was to use this technique to optimize the speed and the accuracy of the identification, as well as to remove the possible ambiguities in the identification obtained by the RFLP of the hsp65 gene. The limited success of this technique can probably be attributed to the taxonomic inflation observed in recent years as well as the polymorphism of the gene coding for 16S rRNA within the Nocardia genus. All of this work constituted a major advance in the identification of species within the Nocardia genus. However, these techniques remain inadequate given the new taxonomic context of the Nocardia genus and novel diagnosis means must be proposed.36 DNA Sequencing. In recent years, the development of novel molecular biology techniques has allowed for the large scale sequencing of DNA. The DNA sequences obtained have been used as taxonomic signatures and allow one to ascribe

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a microorganism to a particular species of taxonomic group. The current taxonomical complexity of the Nocardia genus, which includes over 70 species to date of which at least half are found in clinical settings, requires the use of bioinformatic analysis of ribosomal 16S DNA sequences.37 The combined used of sequences of multiple genes (hsp65, rpoB, sod) should enable one to better discriminate between species in the long term. In fact, the multilocus approach should enable the development of a phylogenetic metatree that should further improve the resolution of the identification within different genera of this family. 16S rRNA Sequencing. To date, the analysis by partial (500 bp) or complete (1500 bp) sequencing of 16S rRNA remains the reference molecular method for the identification of a number of microorganisms. Identification by sequencing is becoming an essential diagnosis tool. Although its cost is limiting for certain clinical biology laboratories, it is a useful alternative for the identification of slow-growing bacteria for which specialized identification techniques are required. However, the precise identification of a microorganism by sequencing will require the availability of sequences in databases open to the general public. Indeed, a monitored database of quality is essential for the successful identification of microorganisms. The use of such a marker for the identification of Nocardia was studied by several authors.30,38–42 The analysis of the complete gene showed that the study of the first 500 base pairs on the 5′ end was insufficient to identify the species of most actinomycetes of medical interest. According to Stackebrandt et al.43 it is now commonly accepted that when the homology between the 16S RNA sequences of two strains is less than 97%, these belong to different species. Several groups have studied its application with the aim to identify bacterial strains within the same species. The work of Mellman et al.40 on a fragment of 500 bp of the 5′ end of the 16S rRNA gene of 30 strains of Nocardia have showed an interspecies polymorphism sufficient for the identification of all tested species, except for N. soli and N. cummidelens. Phenotypic tests were proposed to doubtlessly differentiate these species. The sequences obtained for all these type strains were used to build a genomic bank dedicated to Nocardia. The identification by sequencing of 91 clinical strains originally identified though conventional tests were compared to reference sequences of type strains in the database. The results suggested that a similarity index greater than 99.12% is required to classify a strain as a given species with a statistical probability of error of 1%. Indeed, identification by sequencing of a 500 bp fragment of 16S rRNA of clinical strains yielded excellent results in a majority of isolates. These results were compared with other genomic databases in order to test the degree of accuracy and quality of the identification. With the same goal, Cloud et al.39 studied the identification of Nocardia species of medical interest by sequencing that same 500 bp fragment of 16S rRNA, having chosen a similarity criterion of 99% with a 3% statistical error probability.

Molecular Detection of Human Bacterial Pathogens

Nevertheless, in our work on 5′-end sequencing of the 16S gene of all type strains of the Nocardia genus (43 species described at the time this work was conducted, versus 30 species used in the work of Mellman), we observed the limitations of the use of the gene coding the 16S rRNA as a genotypic marker for the identification of all species. For instance, N. cummidelens and N. soli on the one hand, and N. shimofusensis and N. higoensis on the other, display identical sequences. Similarly, the sequences of N. abscessus and N. asiatica differ by only one nucleotide. Clarridge et al.44 clearly indicate that it is impossible to set a definitive similarity or dissimilarity index for the identification of species by 16S rRNA sequencing, partly because different values are obtained from different databases and using different methods. These facts highlight the need for controlled, monitored, and corrected reference databases. Sequencing of the hsp65 Gene. The hsp65 gene seems to be an interesting target according to studies published on Mycobacterium.45 Rodriguez-Nava et al.42 described a novel molecular technique to differentiate and identify 43 reference species of the Nocardia genus by the amplification and sequencing of a 440 bp fragment of the hsp65 gene. The sequencing of that gene in the Nocardia genus was evaluated through comparison with the sequencing of 16S rRNA. They computed an evolution distances matrix in order to evaluate the interspecies polymorphism of each genetic target and observed an interspecies variability. Results showed sequences similarities on the order of 88%–100% (93.7% on average) for hsp65 on one hand and a 90.5%–100% sequences similarity (94.8% on average) for the gene coding for 16S rRNA. These results demonstrate the utility of hsp65 sequencing: it allows one to differentiate between species having identical 16S rRNA molecules. The study of other genotypic markers among which hsp65, rpoB, recA, sod, and gyrB (used for the identification of mycobacteria) may be considered to solve certain differentiation issues of species within the Nocardia genus. Databases and Biocomputing Tools. The importance of generalized databases is on the rise (EMBL,46 GenBank,47 and DDBJ48). The majority of scientific journals in the molecular biology field systematically require the access number given by the genomic databases when publishing a sequence. Although sequencing methods have been much improved today, the methods used in the 1990s did not always offer quality sequences. Moreover, the lack of quality controls during the submission of sequences to databases inevitably leads to the introduction of erroneous sequences. Specialized databases were therefore created in order to remedy the introduction of errors in general databases. Specialized databases are dedicated to a given biological field. Indeed, they consist of sequences and data that are specific to a given topic. Several specialized genomic databases have been developed in recent years that offer efficient bioinformatic systems for the identification of microorganisms. In the field of actinomycetes, RIDOM, MicroSeq, and, more

Nocardia

recently, BIBI49 databases were developed for Mycobacteria and Nocardia genera. MicroSeq (Microbial Identification System). MicroSeq is a genomic database developed by Perkin Elmer Applied Biosystems. It was created for the identification of filamentous fungi, yeast, and eubacteria. Two kits are commercially available for the identification of bacteria—the MicroSeq500R (which generates a fragment of 500 bp at the 5′ end) and, the most often used, the 16S rRNA (which generates a 1500 bp fragment). MicroSeq contains all reactants and protocols necessary to the sequencing and analysis of results. The database associated with this system includes about 1400 sequences issued from different collections of bacterial strains, notably the most typical strains, including 82 nucleotidic sequences of the 16S rRNA gene of 82 species belonging to the Mycobacterium genus50 and 28 sequences corresponding to 28 species of Nocardia.39 The MicroSeq system of identification functions as follows: an unknown isolate is identified by partial sequencing of the 16S rRNA gene using the MicroSeq kit, and its species is confirmed if the similarity in the sequences is ≥99% as compared to that of a type strain of the MicroSeq database. Regarding Nocardia, Cloud et al.39 tested the MicroSeq system. In order to evaluate the interest of this database, the authors tested 94 strains of clinical origins belonging to the Nocardia genus. The results obtained via sequencing were compared to other molecular studies including restriction enzyme analysis of the gene coding for 16S rRNA or the gene hsp65 and biochemical methods and antibiotic sensitivity. The MicroSeq system was revealed to be capable of correctly identifying only six species that are pathogenic for humans (N. brasiliensis, N. cyriacigeorgica, N. farcinica, N. nova, N. otitidiscaviarum, and N. veterana). However, neither type IV N. abscessus and N. asteroides nor the other species in the genus were clearly identified. One disadvantage of MicroSeq is that it is a closed system. Only bacteria of medical interest have been integrated, which makes it impossible to identify environmental species for which a new implication in human pathology has been observed. In addition, access to the database requires payment. RIDOM (Ribosomal Differentiation of Medical Microorganisms). RIDOM was developed in 1999 by Harmsen et al.51 The aim was to provide bioinformatic services for the identification of microorganisms of medical interest via sequencing. The RIDOM site is directly accessible on the web at http://www.ridom.de. The operation of the intuitive HTML interface of RIDOM is based on two programs: the FASTA program allows one to compare often bulky sequences using a heuristic method; the CLUSTAL W program is used for aligning homologous sequences. The database was constructed using the 16S rRNA sequences of several strain collections of medical interest belonging to several genera including Mycobacterium [in all, 199 sequences of 16S rRNA and 84 sequences of

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internal transcribed spacers (ITS)52] and Nocardia (in all, 30 sequences of 16S rRNA from type strains40). RIDOM’s diagnostic procedure is based on amplifying and sequencing a part of the gene coding for the small subunit of 16S rRNA corresponding to the 5′ region of an unknown specimen. The diagnostic is linked to the analysis of similarities in the sequences sought in the database by the user. The percentage of similarities of sequences is calculated and must be3 99.12% with respect to a type strain in order for a species name to be obtained. In case identification at the species level cannot be provided, the RIDOM system proposes the necessary information to the user that enables him or her to apply a polyphasic approach (combining phenotypical, chemotaxonomic, or molecular approaches). This information is available through hypertext links to relevant online web pages. A study40 sought to evaluate the quality of results provided by two specialized databases (RIDOM and MicroSeq 500) and by the generalized database, GenBank. The results for the three databases were compared with phenotypical identification. This study was carried out via the sequencing of 91 clinical strains using the two systems, RIDOM and MicroSeq separately. A similarity value of 99.12% was set as the identification criterion by these authors. The results obtained using RIDOM showed that among the 91 isolates tested, 65 sequences (71.4%) were identical. On the other hand, 11 isolates (12.1%) showed sequence similarities lower than 99.12%. Meanwhile, the MicroSeq system showed perfect agreement for only 24 isolates (26.4%). Finally, the sequencing results for the ensemble of isolates were compared to those of Genbank. These results suggested that sequences were 100% similar for 54 isolates, although 38 of these isolates are not exclusive to one species. The ensemble of the results obtained through the comparison of these three systems indicates the superiority of RIDOM. Nevertheless, the main disadvantages of RIDOM concerning the Nocardia genus must be noted. First, it is not exhaustive with respect to diagnostic needs (only 30 species are indexed). Second, it is currently impossible to access data concerning Nocardia on the Web site. BIBI (BioInformatic Bacterial Identification). BIBI database aims at facilitating the identification of bacteria, especially in the clinical domain.49 The database uses several bioinformatic tools to automate the analysis of DNA sequences. Two types of databases stem from BIBI: General databases: the major database, “Bacteria,” gathers all the sequences from the GenBank and includes all genes and bacteria species. Specialized and controlled databases: there are five of these databases, corresponding to 16S rRNA, hsp65, rpoB, sod, and gyrB genes. BIBI uses several similar search programs, such as BLAST and CLUSTAL W, applied to the various subsets of sequences from Genbank. These sequences are filtered and stored in a new database designed for bacterial identification.

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The analysis of an unknown sequence is conducted in four steps: search of known sequences similar to the unknown one, extraction of the selected sequences, alignment of the sequence, and display of the results.49 The first step consists of submitting the unknown sequence to the server. The search for similar sequences is conducted through BLAST using a database chosen by the user. The results given by BLAST are then filtered by BIBI and entered in a FASTA file. The unknown sequence and those similar to it are aligned using CLUSTAL W, which performs a multiple alignment, creating three separate files that contain the alignment of sequences, the phylogenetic tree in the NEWICK format, and the distance matrix. The final result is presented in the form of a table of phylogenetic distances between the unknown sequence and all the other similar sequences selected by BLAST. The closest sequences correspond to the identified species. All files produced during this process are accessible online or can be downloaded by FTP “File Transfer Protocol.” BIBI (http://pbil.univ-lyon1.fr/bibi) is a free, simple, and user-friendly bioinformatic tool for the identification of bacteria that quickly gives the user a result while allowing him/ her to monitor the satisfactory progress of every step of the process. As a controlled database, it contains perfect and nonambiguous sequences of normalized size and uses a standardized annotation system. As for the Nocardia and Mycobacterium genera, this specialized database includes all partial sequences of 16S rRNA, hsp65, rpoB, and sod of the type strains of both genera. The Mycobacterium genus contains a total of 388 sequences of 97 species and subspecies that correspond to the four studied genes, 16S rRNA, hsp65, rpoB, and sod.45 The Nocardia genus contains over 200 nucleotidic sequences from over 60 species corresponding to the same four genes.

12.2  METHODS 12.2.1  Sample Preparation Samples can have various origins: bronchoalveolar lavage, aspirations, expectorations, abscess puss, blood (hemocultures are rarely positive, even in the case of disseminated nocardiosis), CSF, pleural liquid, bone tissue, lymphatic ganglions, peritoneal fluid effusion, hepatic biopsy, cutaneous biopsy, pulmonary biopsy, and so forth, and can be collected by puncture, biopsy, during the course of a surgical procedure or even postmortem. In the case of a localized pulmonary infection, bacteria may occasionally be isolated from spital or from material collected through a bronchoscopy, transtracheal aspiration, transcutaneous pulmonary biopsy, or even thoracotomy. However, the presence of Nocardia may only be the consequence of a simple colonization. It is important to note that the enzymatic digestion processes and alkaline decontamination of the expectoration products used for the isolation of mycobacteria alter the viability of Nocardia and greatly reduce the chances of obtaining a culture.

Molecular Detection of Human Bacterial Pathogens

Once pulmonary nocardiosis has been diagnosed, a checkup must always be performed to determining whether the infection has spread and should at least include a cerebral scan, even in the absence of any neurological symptoms. Given cerebral lesions, guided stereotaxic procedures allow for the draining of abscesses and lead to a diagnosis, even for cerebral sites not easily accessible by conventional surgery. Following the diagnosis of an extracerebral nocardiosis, the lesions observed on a scan are presumed to be of the same origin. One must therefore avoid puncturing such lesions for the sole purpose of diagnosis, given the risk of meningitic contamination, except for cases involving AIDS patients for which multiple germ abscesses are common.

12.2.2  Detection Procedures 12.2.2.1  Genus-Specific Identification The work of Laurent et al.27 has enabled the selection of primers within the 3′ region of the ribosomal 16S RNA of Nocardia that allow the specific amplification of a fragment of 596 base pairs for strains belonging to the Nocardia genus. The specificity of the amplified fragment of 596 base pairs can be verified thanks to the position of restriction sites of two enzymes specific to the Nocardia genus: all Nocardia strains tested showed a restriction site specific to the Mnl I enzyme and no restriction site for the enzyme, Sac I. This technique is directly applicable to clinical samples (bronchoalveolar lavage, deep puss, biopsies, CSF or puncture of cerebral abscess) without the need for cell cultures.28 12.2.2.2  Species-Specific Identification Several molecular approaches have been developed for which the reliability has been called into question recently.29,30 The current taxonomical complexity of the Nocardia genus, which includes over 70 species to date of which at least half are found in clinical settings, requires the use of bioinformatic analysis of ribosomal 16S RNA sequences.37 The combined used of sequences of multiple genes (hsp65, rpoB, sod) should enable one to better discriminate between species in the long term. In fact, the multilocus approach should enable the development of a phylogenetic metatree that should further improve the resolution of the identification within different genera of this family.

12.3  CONCLUSIONS AND FUTURE WORK Nocardiosis is an infection which principally affects immunosuppressed patients although immunocompetent subjects may be touched as well. Incidences of Nocardia infection worldwide have been on the rise in recent years, mostly due to the aging of the population and to the usage of aggressive immunosuppressive therapeutics. In France, such infections are often associated with the N. farcinica (26%) and N. nova (20%) species, which are particularly resistant to a number of antibiotics.

Nocardia

Phenotypical techniques used some years ago for the identification of Nocardia were soon replaced by molecular biological methods. These methods have lead to the complete modification of the taxonomy and one can now differentiate more quickly and more accurately between a much larger number of species of the Nocardia genus via specific genetic markers (16S rRNA, rpoB, sod, hsp65). Several difficulties are encountered in the isolation and culture of Nocardia such as their slow growth which can be masked by the presence of other fast-growing microorganisms. To date, this often-tedious procedure can be replaced by a standard real-time PCR reaction carried out directly from a clinical sample without need of culture. Besides, in order to reduce the costs of the analysis and to bring down the time needed for diagnosis to several hours, it would be conceivable the direct detection of Nocardia from a clinical sample or a more complex sample as a soil one by real-time PCR: this could turn out to be of great interest for  the specific and quantitative detection of Nocardia. However the culture must not be neglected because it is necessary for the antibiotic susceptibility testing report. Moreover, other techniques (including mass spectro­ metry), dedicated to the identification of microorganisms through comparison of their protein fingerprints with the aid of a database, may turn out to have interesting implications for the diagnosis of Nocardia.

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1. Boiron, P., and Laurent, F., Nocardia et nocardiose. In Bricaire, L. and Bricaire, F. (Editors), Maladies Infectieuses, pp. 1–8, EMC Elsevier SAS, Paris, 2007. 2. Woese, C.R. et al., The phylogeny of purple bacteria: The alpha subdivision, Syst. Appl. Microbiol., 5, 315, 1984. 3. Stackebrandt, E., Rainey, F.A., and Ward-Rainey, N.L., Proposal for a new hierarchic classification system, Actinobacteria classis nov., Int. J. Syst. Bacteriol., 47, 479, 1997. 4. Lechevalier, H.A., Nocardioforms actinomycetes. In Williams, S.T., Sharpe, M.E., and Holt, J.G. (Editors), Bergey’s Manual of Systematic Bacteriology, pp. 1348–2404, Williams and Wilkins Co, Baltimore, 1989. 5. Chun, J., and Goodfellow, M., A phylogenetic analysis of the genus Nocardia with 16S rRNA gene sequence, Int. J. Syt. Bacteriol., 45, 240, 1995. 6. Goodfellow, M., and Lechevalier, M.P., Genus Nocardia Trevisan 1889, 9AL. In Williams, S.T., Sharpe, M.E., and Holt, J.G. (Editors), Bergey’s Manual of Systematic Bacteriology, vol. 4, pp. 2350–2361, Williams and Wilkins, Baltimore, 1989. 7. Groth, I., and Saiz-Jimenez, C., Actinomycetes in hypogean environments, Geomicrobiol. J., 16, 1, 1999. 8. Schabereiter-Gurtner, C. et al., Phylogenetic 16S rRNA analysis reveals the presence of complex and partly unknown bacterial communities in Tito Bustillo cave, Spain, and on its Palaeolithic paintings, Environ. Microbiol., 4, 392, 2002. 9. Jurado, V. et al., Agromyces salentinus sp. nov. and A. neoliticus sp. nov., Int. J. Syst. Evol. Microbiol., 55, 153, 2005. 10. Beaman, B.L., and Beaman, L., Nocardia species: Hostparasite relationships, Clin. Microbiol. Rev., 8, 213, 1995. 11. McNeil, M.M., and Brown, J.M., The medically important aerobic actinomycetes: Epidemiology and microbiology, Clin. Microbiol. Rev., 7, 357, 1994.

135 12. Exmelin, L. et al., Molecular study of nosocomial nocardiosis outbreak involving heart transplant recipients, J. Clin. Microbiol., 34, 1014, 1996. 13. Beaman, B.L. et al., Nocardial infections in the United States, 1972–1974, J. Infect. Dis., 134, 286, 1976. 14. Saubolle, M.A., and Sussland, D., Nocardiosis: Review of clinical and laboratory experience, J. Clin. Microbiol., 41, 4497, 2003. 15. Boiron, P. et al., Review of nocardial infections in France 1987 to 1990, Eur. J. Clin. Microbiol. Infect. Dis., 11, 709, 1992. 16. Rodriguez-Nava, V. et al., Molecular identification of clinical isolates of Nocardia and related Actinomycetes in the French Observatory for Nocardiosis (OFN)—New epidemiology data of nocardiosis in France between 2000 and 2007. ESCCMID, Abstract number: P1718, Barcelona, Spain, 2008. 17. Rippon, J.W., Nocardiosis. In Rippon, J.W. (Editor), Medical Mycology. The Pathogenic Fungi and the Pathogenic Actinomycetes, 3rd ed., pp. 53–68, WB Saunders Company, Philadelphia, 1988. 18. Filice, G.A., Nocardiosis. In Sarosi, G.A., and Davies, S.F. (Editors), Fungal Diseases of the Lung, 2nd ed., pp. 191–204, Raven Press, New York, 1993. 19. Loeffler, J.M. et al., Nocardial brain abscess: observation of treatment strategies and outcome in Switzerland from 1992 to 1999, Infection, 29, 337, 2001. 20. Laurent, F., Freney, J., and Boiron, P., Nocardia et actinomycètes aérobies apparentés. In Freney, J. et al. (Editors), Actualités en bactériologie clinique théorique et pratique, pp. 1–21, Editions ESKA, Editions Alexandre Lacassagne, 2003. 21. Beaman, B., The pathogenesis of Nocardia. In Fiscetti, V.A., Novick, R.P., Ferretti, J.J., Pottnoy, D.A., and Rood, J.I. (Editors), Gram Positive Pathogens, pp. 594–606, ASM Press, Washington DC, 2000. 22. Boiron, P., Provost, F., and Dupont, B., Technical protocols. In Boiron, P., Provost, F. and Dupont, B. (Editors), Methodes de laboratoire pour le diagnostic de la nocardiose, pp. 107–126, Institut Pasteur, Paris, 1993. 23. Goodfellow, M., The family Nocardiaceae. In Balows, A. et al. (Editors), The Prokaryotes. A Handbook on the Biology of Bacteria: Ecophysiology, Isolation, Identification, Applications, 2nd ed., Vol II, pp. 1188–1213, Springer-Verlag, Berlin, 1992. 24. Wauters, G. et al., Distribution of Nocardia species in clinical samples and their routine rapid identification in the laboratory, J. Clin. Microbiol., 43, 2624, 2005. 25. Lechevalier, H., and Lechevalier, M.P., Classification of aerobic actinomycetes based on their morphology and their chemical composition, Ann. Inst. Pasteur (Paris), 108, 662, 1965. 26. Lechevalier, M.P., and Lechevalier, H.A., Chemical composition as a criterion in the classification of aerobic actinomycetes, Int. J. Syst. Bacteriol., 20, 435, 1970. 27. Laurent, F., Provost, F., and Boiron, P., Rapid identification of clinically relevant Nocardia species to genus level by 16S rRNA gene PCR, J. Clin. Microbiol., 37, 99, 1999. 28. Couble, A. et al., Direct detection of Nocardia spp. in clinical samples by a rapid molecular method, J. Clin. Microbiol., 43, 1921, 2005. 29. Steingrube, V.A. et al., Rapid identification of clinically significant species and taxa of aerobic actinomycetes, including Actinomadura, Gordona, Nocardia, Rhodococcus, Strepto­myces, and Tsukamurella isolates, by DNA amplification and restriction endonuclease analysis, J. Clin. Microbiol., 35, 817, 1997. 30. Conville, P.S. et al., Identification of Nocardia species by restriction endonuclease analysis of an amplified protein of the 16S rRNA gene, J. Clin. Microbiol., 38, 158, 2000.

136 31. Brownell, G.H., and Belcher, K.E., DNA probes for the identification of Nocardia asteroides, J. Clin. Microbiol., 28, 2082, 1990. 32. Laurent, F. et al., Ribotyping: A tool for taxonomy and identification of the Nocardia asteroides complex species, J. Clin. Microbiol., 34, 1079, 1996. 33. Kalpoe, J.S. et al., Molecular typing of a suspected cluster of Nocardia farcinica infections by use of randomly amplified polymorphic DNA, pulsed-field gel electrophoresis, and amplified fragment length polymorphism analyses, J. Clin. Microbiol., 45, 4048, 2007. 34. Steingrube, V.A. et al., DNA amplification restriction endonuclease analysis for differentiation of 12 species and taxa of Nocardia, including recognition of four new taxa within the Nocardia asteroides complex, J. Clin. Microbiol., 33, 3096, 1995. 35. Rodriguez-Nava, V., Devulder, G., and Laurent, F., Taxonomie et identification des Nocardia: évolution et perspectives, Bull. Soc. Fr. Microbiol., 18, 177, 2003. 36. Rodriguez-Nava, V. et al., Nocardia ignorata: A new agent of human nocardiosis isolated from respiratory specimens in Europe and soil sample from Kuwait, J. Clin. Microbiol., 43, 6167, 2005. 37. Patel, J.B. et al., Sequence-based identification of aerobic actinomycetes, J. Clin. Microbiol., 42, 2530, 2004. 38. Cloud, J.L. et al., Identification of Mycobacterium spp. by using a commercial 16S ribosomal DNA sequencing kit and additional sequencing libraries, J. Clin. Microbiol., 40, 400, 2002. 39. Cloud, J.L. et al., Evaluation of partial 16S ribosomal DNA sequencing for identification of Nocardia species by using the MicroSeq 500 system with an expanded database, J. Clin. Microbiol., 42, 578, 2004. 40. Mellmann, A. et al., Evaluation of RIDOM, MicroSeq, and Genbank services in the molecular identification of Nocardia species, Int. J. Med. Microbiol., 293, 359, 2003.

Molecular Detection of Human Bacterial Pathogens 41. Patel, J.B. et al., Sequence-based identification of aerobic actinomycetes, J. Clin. Microbiol., 42, 2530, 2004. 42. Rodríguez-Nava, V. et al., Use of PCR-restriction enzyme pattern analysis and sequencing database for hsp65 gene based identification of Nocardia species, J. Clin. Microbiol., 44, 536, 2006. 43. Stackebrandt, E. et al., Report of the ad hoc committee for the re-evaluation of the species definition in bacteriology, Int. J. Syst. Evol. Microbiol., 52, 1043, 2002. 44. Clarridge, J.E., Impact of 16S rRNA gene sequence analysis for identification of bacteria on clinical microbiology and infectious diseases, Clin. Microbiol. Rev., 17, 840, 2004. 45. Devulder, G., de Montclos, M., and Flandrois, J.P., A multigene approach to phylogenetic analysis using the genus Mycobacterium as a model, Int. J. Syst. Evol. Microbiol., 55, 293, 2005. 46. Baker, W. et al., The EMBL nucleotide sequence database, Nucleic Acids Res., 28, 19, 2000. 47. Benson, D.A. et al., GenBank, Nucleic Acids Res., 28, 15, 2000. 48. Tateno, Y. et al., DNA Data Bank of Japan (DDBJ) in collaboration with mass sequencing teams, Nucleic Acids Res., 28, 24, 2000. 49. Devulder, G. et al., BIBI, a bioinformatics bacterial identification tool, J. Clin. Microbiol., 41, 1785, 2003. 50. Hall, L. et al., Evaluation of the MicroSeq system for identification of mycobacteria by 16S ribosomal DNA sequencing and its integration into a routine clinical mycobacteriology laboratory, J. Clin. Microbiol., 41, 1447, 2003. 51. Harmsen, D. et al., Intuitive hypertext-based molecular identification of micro-organisms, Lancet, 353, 291, 1999. 52. Harmsen, D. et al., RIDOM: Comprehensive and public sequence database for identification of Mycobacterium species, BMC Infect. Dis., 3, 26, 2003.

13 Propionibacterium Andrew McDowell and Sheila Patrick CONTENTS 13.1 Introduction...................................................................................................................................................................... 137 13.1.1 Classification, Morphology, and Biology............................................................................................................. 137 13.1.2 Propionibacteria and Disease............................................................................................................................... 139 13.1.3 Pathogenic Features of Propionibacteria.............................................................................................................. 140 13.1.4 Diagnosis of Human Propionibacterial Infections................................................................................................141 13.1.4.1 Conventional Techniques........................................................................................................................141 13.1.4.2 Molecular Detection...............................................................................................................................143 13.2 Methods............................................................................................................................................................................ 145 13.2.1 Sample Preparation............................................................................................................................................... 145 13.2.1.1 Preparation of Sonicate from Retrieved Medical Devices.................................................................... 145 13.2.1.2 DNA Extraction from Clinical and Bacterial Samples.......................................................................... 145 13.2.1.3 Preparation of Samples for Fluorescent In Situ Hybridization...............................................................147 13.2.2 Detection Procedures.............................................................................................................................................147 13.2.2.1 PCR Detection of Propionibacterium Species.......................................................................................147 13.2.2.2 Nested and Semi-nested PCR Detection of Propionibacterium propionicum and Propionibacterium acnes........................................................................................................................147 13.2.2.3 Fluorescent In Situ Hybridization for the Detection of Propionibacterium acnes............................... 148 13.2.2.4 Nucleotide Sequence Analysis for Differentiation of Propionibacterium acnes Phylotypes............... 148 13.2.2.5 PCR Differentiation of Propionibacterium acnes Types IA, IB, and II............................................... 149 13.3 Conclusions and Future Prespectives............................................................................................................................... 150 Acknowledgments...................................................................................................................................................................... 151 References.................................................................................................................................................................................. 151

13.1  INTRODUCTION 13.1.1  Classification, Morphology, and Biology The genus Propionibacterium is currently composed of the following 11 species: P. freudenreichii (P. freudenreichii subsp. freudenreichii and P. freudenreichii subsp. shermanii), P. jensenii, P. thoenii, P. acidipropionici, P. cyclohexanicum, P. microaerophilum, P. acnes, P. avidum, P. granulosum, P.­ australiense, and P. propionicum. The species Propionibacterium lymphophilum and Propionibacterium innocuum, previously described as part of the Propionibacterium genus, have now been reclassified within the family Propionibacteriaceae as Propionimicrobium lymphophilum and Propioniferax innocua, respectively.1,2 Bacteria within the Propionibac­terium genus are gram-positive, anaerobic-to-aerotolerant-to-microaerophilic, nonspore-forming pleomorphic rods. They are 0.2–1.5 μm in diameter × 1–5 μm in length and have a high G + C content [57–68 mol% (Tm)]. They can appear diphtheroid or club shaped, with one end rounded and the other tapered or pointed, but may also be coccoid, bifid, branched, or filamentous with lengths of up to 20 μm.3 Propionibacteria are chemoorganotrophs with

complex nutritional requirements and produce large amounts of propionic and acetic acids, and lesser amounts of isovaleric, formic, succinic or lactic acids, and carbon dioxide as the end products of their fermentation reactions.3 Although predominantly anaerobic, most propionibacteria are catalase positive. Tetrahydrogenated menaquinones with nine isoprene units [MK-9(H4)] are the major respiratory quinones. Their optimal growth temperature is between 30°C and 37°C, and on solid media colonies will appear smooth, rough, or convex, and may be white, grey, pink, red, yellow, or orange in color.3 Within the genus, species have been traditionally differentiated into two groups, the “classical” or “dairy” and the “cutaneous” propionibacteria. While molecular analyses have revealed that the phylogenetic relationships amongst propionibacteria do not exactly mirror their classification based on habitat (Figure 13.1), it is still a useful and convenient way to group the species. The “classical” or “dairy” propionibacteria includes the species P. freudenreichii, P.  jensenii, P.  thoenii, and P. acidipropionici, which are isolated mainly from cheese and other dairy products and are also used as starter cultures by the dairy industry and 137

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Molecular Detection of Human Bacterial Pathogens

P. acidipropionici (ATCC 25562)

95

P. microaerophilum (DSM 13435)

99

P. jensenii (ATCC 4868) 100

100

P. thoenii (ATCC 4874) P. granulosum (DSM 20700)

91

P. acnes (ATCC 6919) P. avidum (ATCC 25577)

100 100

P. propionicum (ATCC 14157) P. cyclohexanicum (ATCC 700429)

90

P. australiense (ATCCBAA-264) 100

P. freudenreichii subsp. freudenreichii (ATCC 6207) 100

P. freudenreichii subsp. shermanii (ATCC 9614) N. asteroides (ATCC 19247)

0.01

FIGURE 13.1  Phylogenetic tree of 16S rRNA sequences for the genus Propionibacterium. The neighbor-joining tree was constructed using the Jukes-Cantor-based algorithm. The sequence input order was randomized, and bootstrapping resampling statistics were performed using 100 data sets. The 16S rRNA sequence from Nocardia asteroides was used as a distant outgroup to root the tree as it also belongs to the order Actinomycetales. Bootstrap values are shown at each node of the tree.

as protective biopreservatives and probiotics.4 These organisms have also been isolated from silage, fermenting olives, and soil.5–8 The “cutaneous” propionibacteria, which include the species P. acnes, P.  granulosum, and P. avidum, are isolated from human skin, although they also colonize the mouth, upper respiratory tract, large intestine, and genitourinary tract.3,9 There have also been reports in the literature describing their isolation from animal species such as cows, pigs, and dogs.10–12 Levels of P. acnes colonization on the skin vary from person to person and from the region of the body sampled.13 P. acnes is especially prevalent in sebaceous gland-rich areas on the skin, with the neck, forehead, and shoulder showing some of the highest concentrations compared to other sites, such as the abdomen, hip, knee, and chest where levels are lower.14 Propionibacterium granulosum is present on the sebum-rich, oily areas of the skin, and is especially common in the ala nasi (wing of the nose), although the total numbers present are much less than P. acnes.15 The distribution of P. avidum appears more restricted and is found principally in moist areas, including vestibule of the nose, the axilla, and the perineum.15 The “cutaneous” group has also been known as the “anaerobic coryneforms,” “anaerobic diphtheroids,” “acnes group strains,” or “cutaneous propionbacters.” The more recently described Propionibacterium

species, P. microaerophilum, P. cyclohexanicum, and P. australiense, do not conveniently fit into either the “classical” or the “cutaneous” groups based on habitat.16–18 On the basis of 16S rRNA sequence, however, P. microaerophilum, recovered from a decantation reservoir of olive mill wastewater, clusters with the “dairy” propionibacteria, while P. cyclohexanicum and P. australiense, recovered from spoiled orange juice and granulomatous bovine lesions, respectively, cluster with one another and are more closely related to P. freudenreichii than any of the other propionibacteria.16–18 The species P. propionium, previously known as Arachnia propionica, clusters with the “cutaneous” species based on 16S rRNA sequence and is present in the oral cavity of humans including the mucosa, tonsils, and dental plaque, and may also be present in the oral cavity of other animals.19 It has also been isolated from the gastrointestinal and female genital tract, but it is not believed to be part of the indigenous microbiotia in these areas.19 Morphologically, the “classical” propionibacteria tend to be shorter and thicker than the “cutaneous” group, which are more often irregular and curved, although all the strains may be variable in morphology, especially in early exponential phase cultures.3 Strains of P. acnes can give rise to longer and more slender irregular rods in young cultures,

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Propionibacterium

while in stationary phase cultures all strains tend to be more coccoid.3 The cell dimensions of P. acnes range from 0.5 to 1.5 µm in width and approximately 1–5 µm in length.20 Propionibacterium propionicum and strains of P. acnes phylotype III can form filaments.21,22 With P. propionicum, these filaments are branched and are especially evident in young liquid media cultures, but commonly break up into short rods in older cultures. Morphologically and phenotypically, P. propionicum resembles Actinomyces israelii. Some strains of propionibacteria (e.g., P. thoenii; P. avidum) appear capsulated, and it seems that a considerable number of strains of all species may produce extracellular slime not organized in the form of clear-cut individual capsules.23,24 The extracellular material is carbohydrate in nature.23,25 The cell lipids of both “classical” and “cutaneous” strains contain large amounts of C15 branched-chain fatty acids.26 Both groups of propionibacteria tend to have similar nutritional needs, including a requirement for pantothenate. Many also need biotin and some require thiamine and p-aminobenzoic acid. For others, thiamine and p-aminobenzoic acid are not essential but do stimulate growth, as does nicotinamide and oleate.27–29 In contrast, P. microaerophilum does not require any of these growth factors and is therefore prototrophic. Amino acid requirements are complex for the “cutaneous” group.28 Most strains of P. acnes and P. granulosum require a full complement of 18 amino acids for good growth, which can be enhanced further by the addition of lactate, pyruvate, ketoglutarate, as well as guanine and adenine (20/1 ratio).28 Some strains may also require additional factors, such as heme and vitamin K. In contrast, after several transfers P. avidum can grow in a simple medium containing salts, glucose, and vitamins in the absence of amino acids.28 Propionibacterium acnes has been grouped into three distinct clades, known as phylotypes I, II, and III, based on the analysis of the recA housekeeping gene and a more variable hemolysin/cytotoxin gene.22,30 While the term phylotype has a somewhat arbitrary definition, we have used it in a straightforward context to describe the previous P. acnes types resolved phylogenetically. The phylotype I grouping can also be further subdivided into two very closely related clusters known as IA (including type strain ATCC6909) and IB. While these different lineages can be identified upon sequence analysis of recA, they share at least 99.8% identity in their 16S rRNA sequences (1484 bp section).22,30 Earlier studies by Johnson and Cummins demonstrated that strains now known to represent P. acnes phylotypes I and II share DNA–DNA hybridization values of 88%–99%.31 Combined with more recent 16S rRNA sequence data, this demonstrates that these lineages represent distinct sequence clusters within P. acnes and not novel species, at least based on current definitions of a bacterial species.32,33 The phylotypes of P. acnes can also be distinguished by serological agglutination tests and monoclonal antibody-labeling methods, which reflect differences in the composition of their cell walls.22,30,31 Previous work has shown that phylotype I

strains contain glucose, mannose, and galactose in their cell wall, while phylotype II strains contain only glucose and mannose.31 Data on the composition of type III cell walls is currently not available. While types I and II both contain LL-diaminopimelic acid (LL-DAP) in their cell wall peptidoglycan, some strains of phylotype II may occasionally contain meso-DAP. Other phenotypic differences between these groupings include variation in their susceptibility to bacteriophage infection and distinct fermentation profiles with sugar and sugar alcohols.34,35 The “cutaneous” species P. avidum can also be split into two distinct serovars.31,36 By DNA–DNA hybridization, sequences within each group are at least 90% similar, but between the two groups the average similarity is 80%.37 The cell wall composition of each group mirrors that found in P. acnes phylotypes I and II, with P. avidum type I containing glucose, mannose, galactose, and LL-DAP, while phylotype II contains glucose, mannose, and mostly meso-DAP, but sometimes LL-DAP. Strong cross reaction between P. acnes phylotype II strains and sera to P. avidum type II has been observed.31,36 To date, phylogenetic differences between the P. avidum serovars have not been investigated. In contrast to both P. acnes and P. avidum species, P. granulosum appears to belong to a single serological type.36 Two serovars of P. propionicum have been identified using fluorescent antibody methods with the type strain ATCC14157, representing serovar 1.38,39 No cross reaction between the serovars has been observed by cell wall agglutination tests, but some reaction between serovar 1 and P. avidum has been observed by immunofluorescence.38 The two serovars of P. propionicum share very little DNA homology (1%) and also form distinct subclusters by numerical phonetic analyses suggesting they may actually represent distinct species.40

13.1.2  Propionibacteria and Disease Within the Propionibacterium genus, the “classical” or “dairy” propionibacteria are nonpathogenic, but strains of the “cutaneous” species and P. propionicum are associated with opportunistic human infection and disease.4 Propionibacterium acnes represents the major opportunistic human pathogen within the “cutaneous” group and although P. granulosum has been found in sarcoid tissue and associated with acne; endocarditis, endophthalmitis, and septicemia in an immunocompromised patient; and P. avidum with septicemia and severe infection after invasive procedures; they are still considered a relatively rare cause of human disease based on literature reports (although more cases of P. granulosum infections have been described compared to P. avidum).41–47 Propionibacterium australiense has been shown to cause granulomatous lesions in cattle from different geographical sites in northern and central Queensland, Australia.48 These lesions are present in large numbers throughout the animal, including the external surface of the rumen and the reticulum, tongue, and peritoneal surface of the gastrointestinal tract. The lesions consist of a characteristic fibrous outer capsule that surrounds thick yellow viscous pus with caseating

140

granulomas. To date, there are no reports of this organism causing human infections. Since both P. acnes and P. propionicum represent the major human pathogens within the Propionibacterium genus, their association with infection and disease will now be described in more detail. Propionibacterium acnes is a widespread opportunist most noted for its potential involvement in the inflammatory condition acne. It is also a cause of biofilm-associated infections of indwelling medical devices, such as prosthetic hip, knee, and shoulder joints, spinal implants, prosthetic heart valves, intravascular catheters, and central nervous system shunts.49–58 Infection of orthopaedic implants with P. acnes can be insidious with delayed presentation of symptoms; indeed, many of the traditional signs of underlying infection are absent.52,55 It can, however, also cause acute infection associated with severe tissue damage and sinus tract formation.59 The bacterium is also responsible for endophthalmitis posteye surgery and has been linked to synovitis acne pustulosis hyperostosis osteitis (SAPHO) syndrome, a chronic condition that describes an association of characteristic bone, joint, and skin lesions, and sarcoidosis, a multisystem granulomatous inflammatory disease that is characterized by noncaseating granulomas.60–62 The bacterium may also have a role to play in primary biliary cirrhosis and the development of prostate cancer via chronic low-grade infection of the prostate gland.63,64 In addition to its role in human infection, P. acnes has recently been identified as the cause of necrosuppurative placentitis and abortion in an adult Holstein cow.10 An emerging number of studies have reported the association between the phylotypes of P. acnes and different infections. Studies by Sampedro et al. found that the total distribution of phylotypes I (collectively IA & IB) or II between prosthetic hip, knee, shoulder, and spine implants, removed because of aseptic failure or underlying infection, was similar, although all isolates recovered from knee, shoulder, and spine prostheses were predominately phylotype I (IA & IB), with equal numbers of types I and II being isolated from prosthetic hips.54 They did show, however, that within the phylotype I grouping, isolates of type IB were much more commonly associated with the implant infection cohort compared with aseptic failure, where equal number of types IA and IB were found. The isolation of P.  acnes in association with implants that fail aseptically may reflect colonization of the device prior to the development of infection.51 Studies of P. acnes isolates recovered from the forehead skin of healthy volunteers versus those isolated from infected tissue adjacent to orthopaedic implants found the distribution of P. acnes phylotypes to be similar, although type IA was the predominant phylotype isolated.65 Studies by Shannon et al. found that phylotypes IB and II were more frequently isolated from the radical prostatectomy specimens of patients with prostate cancer, although no significant difference in the prevalence of these P. acnes types in the urinary tract of patients with prostate pathology compared with normal control men was observed.66 Types IB and II were, however, more common in adult men compared with adolescents suggesting an age-related increase.66

Molecular Detection of Human Bacterial Pathogens

Propionibacterium propionicum causes human infections that may be indistinguishable from Actinomyces species based on clinical symptoms and pathogenesis. The bacterium is a recognized cause of actinomycosis, a disease that is characterized by a chronic and subacute granulomatous infection that commonly affects the cervicofacial area (face and neck), but can also affect the thorax, abdomen, or pelvis.67 Aggressive long-term, high-dose, antibiotic therapy is required and if left untreated the infection may spread to the brain or other organs to become life threatening. It is also a common cause of lacrimal canaliculitis, a disease where the tear duct becomes blocked with the bacterium and concomitant oral or nasal microbiota.68 This condition presents mainly in females over 40 years old and can respond poorly to topical antimicrobial treatment. The organism is also associated with primary and persistent ondodontic infection as well  as intrauterine contraceptive devices and has been described as the cause of a finger infection following a human bite, and as a cause of a renal abcess.67,69,70

13.1.3  Pathogenic Features of Propionibacteria Previous studies indicate that strains of P. granulosum have the ability to produce active deoxyribonuclease, lecithinase, hyaluronidase, and a lipase that is considerably more active than that of P. acnes.71–73 The organism does not, however, produce gelatinase and it is also considered to be nonhaemolytic.3 In contrast, P. avidum strains do produce hemolytic and gelatinase activities as well as deoxyribonuclease and proteinase. Most are, however, negative for lecithinase, hyaluronidase, and chondroitin sulfatase.71,72 Propionibacterium acnes also produces extracellular enzymes involved in tissue degradation, including ribonuclease, neuraminidase, hyaluronidase, acid phosphatase, lecithinase, lipase, and proteinase.71,74–80 More detailed insights into the pathogenic potential of P. acnes have been determined by analysis of the whole genome sequence of the phylotype IB strain KPA171202 and the draft genome sequence of the phylotype IA strain ATCC6919.81,82 This has revealed a gene family consisting of five homologues of the Streptococcus agalactiae cohemolysin or CAMP (Christie Atkins Munch Peterson) factor. The gene family is present in all P. acnes phylogenetic groupings described to date and the cohemolytic reaction, a synergistic lysis of erythrocytes by the interaction of the CAMP factor with Staphylococcus aureus sphingomyelinase C, has been observed amongst isolates representing types IA, IB, and II.82 The CAMP reaction in phylotype III strains has not been examined. Differential production of the different CAMP factor proteins amongst P. acnes phylotypes has been found, with strains of phylotype II and a commonly isolated phylotype IB strain (ST-10 by MLST; McDowell & Patrick, unpublished) producing large quantities of CAMP factor 1 compared to phylotype IA organisms.82 In addition, CAMP factor proteins have been shown to bind the Fc fragment of immunoglobulins of the IgG and IgM class and also to act as pore-forming toxins.83 They are also lethal when administered to rabbits and mice.84 A number of putative hemolysin

141

Propionibacterium

genes have also been identified within the genome sequences, including homologues of tlyC and tlyA.30,81 Variable hemolysis of phylotype I strains on horse blood has been observed, but phylotype II and phylotype III strains are nonhemolytic under these conditions.22 Genes coding for surface proteins with cell adherent properties have also been described. These proteins contain the C-terminal–LPXTG—cell wall sorting signal that is acted on by the transpeptidase sortase. Three genes have also been described that contain contiguous stretches of 12–16 guanine or cytosine residues within the promoter or 5′ end regions. These homopolymeric tracts may cause slipped strand mispairing during replication, leading to the expression of proteins with altered molecular masses, thus leading to phenotypic variation and possibly immune evasion or adaptation to environmental changes.85 Phase variation due to slipped strand mispairing has been observed in several notable pathogens (e.g., Neisseria meningitis).86 The strong proinflammatory nature of P. acnes has been known for some time. Indeed, the immunostimulatory activity and adjuvant properties of P. acnes isolates previously referred to as “Corynebacterium parvum” are exploited in studies of the inflammatory response.87 Furthermore, P. acnes has also been used as a nonspecific immune stimulant in the equine industry (EqStim®).88 It can activate complement by both the classical and alternate pathways leading to inflammatory processes through antibody-dependent and independent mechanisms.89 Homologues of GroEL, DnaK, and several other heat shock proteins, which are major targets for the immune system, are also present in the genome.81,90 Three gene clusters putatively involved in extracellular polysaccharide/lipoglycan biosynthesis have also been identified in the genome sequence and may also be important in modulating immunogenicity. The production of porphyrins by P. acnes may lead to inflammation as well as generate toxic reduced oxygen species that cause cell and tissue damage. It has been suggested that porphyrin production by the bacterium may also be important in the skin condition progressive macular hypomelanosis.91 Studies in vitro have shown that P. acnes can stimulate the production of antimicrobial peptide and proinflammatory mediators from human keratinocytes and sebocytes and differences have also been noted in the capacity of strains representing phylotypes IA and IB to modulate the viability and differentiation of these cell types.92 However, further studies with a larger number of strains will be required to demonstrate that these differences are truly phylotype-specific. The production of biofilm is a well-established virulence determinant used by pathogenic bacteria, and the formation of biofilm by P. acnes has been demonstrated by various studies, although the underlying molecular details of biofilm regulation in P. acnes have not yet been determined.93–94 It has been shown in vitro that isolates associated with orthopaedic implant infections produce greater levels of biofilm when compared to isolates from the skin of healthy volunteers.65 Furthermore, P. acnes within a biofilm is not only more resistant to antibiotics, but the production of lipase by the bacterium is upregulated by this mode of growth.94 P. acnes is also resistant to killing and

degradation by human phagocytic cells and can thus survive intracellularly. It may, therefore, persist and cause a chronic low-grade infection. Despite its pathogenic potential, P. propionicum appears unremarkable in its production of specific virulence factors.3 Strains cause β-hemolysis on human blood agar and are variable in the production of lipase. No hyaluronidase, deoxyribonuclease, or proteolytic activity is produced, although some strains may liquefy gelatin. A study of P. acnes isolates causing surgical, foreign body, and septicemia infections in Europe has found that 3% are resistant to tetracycline, 15% to clindamycin, and 17% to erythromycin, while none are resistant to linezolid, benzylpenicillin, or vancomycin.95 The use of topical formulations of erythromycin and clindamycin to treat acne has resulted in cross-resistant propionibacteria strains in Europe and Asia.96– 98 Strains of P. acnes are resistant to metronidazole, sulfonamides, and aminoglycosides.99–101 The current antimicrobial susceptibility of P. granulosum isolates has not been as widely investigated as P. acnes, but studies with two isolates responsible for septicemia and endocarditis in compromised patients revealed susceptibility to a range of compounds.42,44 The septicemia isolate was sensitive to penicillin G, ampicillin, erythromycin, cefotaxime, clindamycin, rifampin, and imipenem, but resistance was found to metronidazole, sulphonamide, fosfomycin, and tobramycin.44 Gentamicin, streptomycin, amikacin, pefloxacin, and chloramphenicol had poor activity. This data appeared to substantiate previous studies.102 The endocarditis isolate was susceptible to penicillin, clindamycin, augmentin, piperacillin, chloramphenicol, cefotaxime, and cefazolin, but resistant to metronidazole.42 Erythromycin-resistant strains of P. granulosum have been isolated from the skin of acne patients treated with oral erythromycin and topical clindamycin.103 Little information is available on the current susceptibility of P. avidum isolates. Susceptibility testing of a P. avidum isolate responsible for a breast abscess following reduction surgery revealed sensitivity to penicillin G, ampicillin, erythromycin, tetracycline, and vancomycin, but resistance to metronidazole.47 Much earlier studies by Hoeffler et al. found that isolates were generally sensitive to a range of drugs including benzylpenicillin, ampicillin, cephalothin, rifampin, clindamycin, erythromycin, and minocycline.102 As with other “cutaneous” propionibacteria, erythromycin-resistant strains have been isolated from the skin of acne patients treated with oral erythromycin and topical clindamycin.103 Propionibacterium propionicum appears sensitive to a wide range of antibiotics such as penicillins, cephalosporins, chloramphenicol, tetracyclines, erythromycin, clindamycin, and imipenem, but fluoroquinolones, aztreonam, aminoglycosides, and metronidazole are ineffective.67

13.1.4  Diagnosis of Human Propionibacterial Infections 13.1.4.1  Conventional Techniques Isolation of Propionibacterium Species from Clinical Samples. Good growth of all propionibacteria can be achieved

142

Molecular Detection of Human Bacterial Pathogens

using a medium with the following composition; trypticase (1% w/v), yeast extract (0.5% w/v), glucose (1% w/v), CaCl2 (0.002% w/v), MnSO4 (0.002% w/v), NaCl2 (0.002% w/v), K-phosphate buffer (0.05 M), Tween-80 (0.05% w/v), sodium dithionite (0.05% w/v), NaHCO3 (0.1% w/v; added as sterile solution along with inoculum), and pH 7.0.104 Complex media, such as brain heart infusion broth or agar, is often used for the growth of “cutaneous” strains which will also grow very well on anaerobic blood agar. A medium containing sodium oleinate for the growth of propionibacteria and inhibition of Actinomyces strains from human skin swabs has also been described.105 The addition of bromocresol purple to the medium permits identification of P. acnes, which appears as yellow or slightly yellow opaque colonies with yellow zones of lysis due to the fermentation of glycerol, while Staphylococcus epidermidis appears as white translucent colonies. Propionibacteria can also be recovered using TYEG agar (2% w/v tryptone, 1% w/v yeast extract, 0.5% w/v glucose) containing furazolidone (6 mg/ml) to inhibit staphylococcal growth.96 Although some strains of propionibacteria may be aerotolerant, a culture methodology similar to that used for the primary isolation for strict anaerobes should still be adopted, such as that detailed in the Wadsworth KTL Anaerobic Bacteriology Manual.106 It is important that growth media and diluents are pre-reduced and pre-equilibrated under anaerobic conditions. Furthermore, the incorporation of l-cysteine hydrochloride (0.05% w/v) into liquid media and diluents provides a suitable reducing agent. An anaerobic gas atmosphere within an anaerobic jar or cabinet, from which oxygen has been scavenged by palladium catalysts, should be used for culture. A lack of adherence to strict anaerobiosis when handling and processing clinical specimens will result in reduced detection of pathogenic Propionibacterium species.49,57 Furthermore, as P. acnes causes biofilm-associated infection of prosthetic implants, it is imperative that removed devices undergo sonication to detach any adherent bacteria.49,50,53 Failure to do so will result in poor diagnostic sensitivity. It is worth highlighting

that sonication is superior to alternate “scraping” methods for the detachment of bacteria growing on titanium and steel surfaces in vitro.107 The “cutaneous” group strains grow at 36°C–37°C, in contrast to the “classical” propionibacteria, which grow best at 30°C–32°C. In general, while maximum growth in complex media of pure cultures is attained in 48 h, primary isolation from clinical samples may require incubation for 7–14 days. Biochemical Analyses. The “cutaneous” propionibacteria and P. propionicum can be differentiated by using a number of simple biochemical tests (Table 13.1). These include sucrose and maltose fermentation, gelatin liquefaction, esculin and casein hydrolysis, and catalase, indole, and nitrate production. A simple thin-layer chromatography (TLC) method has also been described for the differentiation of P. acnes from P. propionicum based on orcinol glycolipid profiles.108 A number of studies have evaluated commercial phenotypic identification kits for their ability to differentiate “cutaneous” propionibacteria, although the number of species and strains examined were limited. The RapID™ ANA II System (Remel, Thermo Fisher Scientific) correctly identified 21/23 P. acnes strains (91%), while the Rapid ID 32A system (bioMérieux) was able to correctly identify all 10 P. acnes strains (100%) under investigation.109,110 Similarly, the BBLCrystal™ Anaerobe (ANR) identification system (Benton Dickson) correctly identified all P. acnes (n = 10), P. avidum (n = 9), and P. granulosum (n = 7) isolates analysed.111 In contrast, problems have been encountered with the API (RAPID) Coryne system and database 2.0, which covered 49 taxa.112 Of six P. acnes strains tested, only one was correctly identified with additional tests, while four were not identified, and one was misidentified. With four P. avidum strains, two were correctly identified, and two could only be identified with additional investigations. Within P. acnes, phylotype I strains can be identified from those of type II and III based on the fermentation of sorbitol and also clear zones of β-hemolytic activity on anaerobic blood agar plates. While these simple tests

TABLE 13.1 Biochemical Tests to Differentiate Propionibacterium Species Associated with Human Infectiona Characteristic Sucrose fermentation Maltose fermentation Gelatin hydrolysis Esculin hydrolysis Casein hydrolysis Catalase production Indole productionb Nitrate reductionb

P. acnes

P. granulosum

P. avidum

− − + − + d+ d+ d+

+ d+ d− − − + − −

+ + − + + + − −

P. propionicum + + − − + − − +

 Symbols: +, positive reaction in 90% or more of isolates; −, negative reaction in 90% or more of isolates; d+, positive reaction in 40%–90% of isolates; d −,

a

positive reaction in 10%–40% of isolates.

 Indole production and nitrate reduction are the main characteristics used to differentiate P. acnes and while the majority of isolates are positive for these

b

reactions, it is important to be aware that isolates negative for these tests do exist. For example, Sampedro et al. found that 15% of P. acnes isolates were negative for indole production.54

Propionibacterium

are 100% specific for type I organisms, they are not 100% sensitive. Types I and II can also be differentiated from each other based on cell wall sugar composition.31 As highlighted in Section 13.1.1, the serotypes of P. acnes, P. avidum, and P. propionicum can be differentiated using cell wall agglutination and fluorescent antibody-labeling methods. 13.1.4.2  Molecular Detection Molecular-based detection methods for bacterial infections have a number of notable advantages over traditional culture-based methods, including increased sensitivity and the opportunity to detect viable but nonculturable (VBNC) bacteria that arise due to stress or unfavorable growth conditions. Although dormant, such organisms may retain their pathogenicity and have the capacity to “resuscitate” and initiate infection.113 In addition, components of nonculturable bacteria may drive an inflammatory response. Bacteria can enter the VBNC state when growing as a biofilm, which may impact on the cultivation of medical device-related infections. The administration of prophylactic antibiotics prior to surgery may also interfere with the growth of pathogenic species in tissue specimens giving rise to false-negative culture reports. Antibiotic challenge may also induce a dormant state akin to VBNC cells.114 Another significant advantage of a molecular approach is the shorter time between receipt of a clinical sample and confirmation of infection, leading to earlier antimicrobial intervention and patient management/ care. This is especially important when dealing with organisms, such as P. acnes, which can require extended anaerobic growth conditions, or infections due to Mycobacterium tuberculosis, which can be slow and cumbersome to detect using conventional culture methods.115 Identification of bacteria based on phenotypic properties can also be unstable at times and dependent upon changes in environmental conditions such as growth substrate, temperature, and pH levels.116 Molecular-based detection of propionibacteria in clinical samples via conventional or real-time PCR amplification of the small subunit ribosomal RNA (16S rRNA) gene has been the predominant approach described in the literature (Table 13.2). Studies have utilized either species-specific 16S rRNA primers to directly target the bacteria or have used universal or broad-range primers that amplify all bacterial sequences present in the sample, followed by nested PCR with “internal” species-specific 16S rRNA primers or, more commonly, separation of the mixed amplicons by cloning and identification by comparison of the nucleotide sequences to those present in online databases, such as GenBank and the Ribosomal Database project.119,125,129–131 While broad-range 16S rRNA PCR can facilitate the identification of multiple species causing polymicrobial infections, the method can be challenging since bacterial or DNA contamination can occur at any stage in the protocol, potentially leading to false-positive results. Sources of contamination range from surgical or laboratory personnel, nonsterile sites surrounding the clinical specimen, the specimen container, DNA extraction and PCR setup reagents and vials, pipettes, PCR cabinets, and aerosols.132 Contamination may also occur due to “carryover” amplicons

143

from previous PCR experiments, particularly when using a nested-PCR methodology. To avoid such problems, a range of preventative measures are required including the use of molecular grade reagents, protective clothing, sterile gloves, rooms specifically designated for DNA extraction and PCR setup, and containing equipment and reagents that must stay on site, filtered tips, HEPA-filtered safety cabinets with UV light for DNA degradation, chemical decontamination of work areas, incorporation of dUTP into PCR reactions and enzymatic treatment of new samples with Uracil-DNAglycosylase, and the addition of appropriate control samples. Scrupulous adherence to aseptic processing protocols and the inclusion of multiple negative control checks is particularly important for the detection of “cutaneous” propionibacteria in samples, even when adopting a species-specific rather than broad-range PCR methodology. While not conclusive, such an approach will provide some reassurance that the organisms detected are not contaminants. In addition, if Propionibacterium species are repeatedly isolated from an infection, this should provide very strong evidence that the bacterium is not a contaminant. For definitive evidence of propionibacterial infection, however, a polyphasic approach should be adopted where possible, including parallel culture experiments on agar plates (to assess bacterial counts) as well as other nonculture-based detection methods, such as Gram staining, immunofluorescence microscopy (IFM), and/or in situ hybridization (ISH). Furthermore, the potential presence of PCR inhibitors in clinical samples also means that internal amplification controls targeting human genes, such as the β-globulin gene, should also be included to rule out false-negative results. In addition to PCR, ISH-based methodologies have also been used for the molecular identification of P. acnes in prostate and sarcoid tissue and to screen isolates from human blood cultures (Table 13.2). This method, which normally uses a single oligonucleotide probe (15–30 nucleotides) that is covalently linked to an enzymatic (e.g., horseradish peroxidase) or fluorescent (e.g., fluorescein isothiocyanate) reporter molecule (FISH), targets rRNA sequences within the bacterial ribosome.133 As with IFM, it provides the opportunity to directly observe, in situ, microbial cells in a sample, and when used with confocal laser scanning microscopy can provide information on the morphology and spatial arrangement of infecting organisms.134 Inherent problems can arise, however, including autofluorescence from tissue samples, photobleaching, insufficient penetration of the probe into the bacterial cell, and weak signals due to low ribosome number. The latter can be a particular problem with slow-growing or dormant organisms. Inaccessibility of the target sequence for hybridization can also occur due to the formation of hairpin loops, and poor signal intensities may further result from self-annealing and hairpin formation of the probe itself.135 In addition to detecting propionibacteria in clinical samples, molecular methods have also been used to characterize isolated organisms (Table 13.2). Differentiation of P. acnes isolates from other “cutaneous” propionibacteria has been achieved by PCR targeting of the 16S rRNA and lipase gene

16S rRNA; Nt-PCR 16S rRNA; RT–PCR 16S rRNA; ISH 16S rRNA; SNt-PCR

16S rRNA; SNt-PCR Lipase; SNt-PCR

16S rRNA; RT–PCR 16S rRNA; FISH 16S rRNA; RT–PCR 16S rRNA; RT–PCR 16S rRNA; PCR 16S rRNA; PCR 16S rRNA; PCR 23S rRNA; FISH

Gene/Methoda

CCCTGCTTTTGTGGGGTG CGCCTGTGACGAAGCGTG CGACCCCAAAAGAGGGAC TCACTGATGAAGATCAACGCAC ACTTCTCCTGCAAGGTCAAG TGCGAATGTCCGACGAAGTCGA GACGGTAGCAGTAGAAGAAGCAC CTGTAAACCGACCAAAAAGG GCGTGAGTGACGGTAATGGGTA TTCCGACGCGATCAACCA ATGGATCGGGAGTGCTCAGAGATGGG AAGGCCCTGCTTTTGTGG TCCATCCGCAACCGCCGAA ACTCACGCTTCGTCACAG

AACCGCTTTCGCCTGTGA ACGCTCAGGGTTAAGCCC ACATGGATCCGGGAGCTTC ACCCAACATCTCACGACACG GGGTTGTAAACCGCTTTCGCCT GGCACACCCATCTCTGAGCAC GGAAACCTGATGCTTAACGT GGCACCAACCATCTCTGGAA GGGTTGTAA(A/T)CCGCTTTCGCCTG GGGACACCCATCTCTGAGCAC GAGTGTGTGAACCGATCATGTAGTAGGCAA

Pa-F Pa-R Pg-F Pg-R PA-F’ PA-R PG-1 PG-2 PA-F′Mb PA-RM PAC23Sc

PAS9 PAS10 PAS11 Lip1 Lip2 Lip4 PP-1 PP-2 PAC-F PAC-R PAC16S Pa1 rPa2 rPa3

AAGCGTGAGTGACGGTAATGGGTA CCACCATAACGTGCTGGCAACAGT GCCCCAAGATTACACTTCCG

Sequences (5′–3′)

PArA-1 PArA-2 PAC16S598

Primers/ Probes

567f 131 − 387/160g

60 42 62

946/573d 515/277e

−

587 426 587

55

75°C followed by 37°C 57 57

54 58 50

182 102

–

46 62 60

677

Amplicon Size (bp)

55–62

Anneal/Hybrid Temperature (°C)

Vitreous fluid; Aqueous humour

Isolates; Root canal samples Sarcoid lymph tissue Sarcoid lymph tissue

Isolates Isolates

Prostate tissue

Prostate; Sarcoid lymph/lung tissue; Vitreous fluid Sarcoid lymph/lung tissue; Vitreous fluid Liver tissue

Bronchoalveolar Lavage cells

Isolates

Prosthetic shoulder sonicate; isolates

Sample(s) Analyzed

127,128

126

69,125

124 124

41,119 120,121 122 41,120 121 63 123

118

117

53,54

Reference

b 

a 

RT–PCR = Real-time PCR; FISH = Fluorescent in situ hybridization; SNt-PCR = Semi-nested PCR; Nt-PCR = Nested-PCR; ISH = In situ hybridization. Primers PA-F'M and PA-RM are modification of primers PA-F' AND PA-R.120 c  FISH probe PAC23S used with four probes generated from P. acnes strain KPA171202 by PCR and cloning (see Section 2.2.3). d  Amplicon sizes of 946/573 bp correspond to first round (PAS9/PAS11) and second round (PAS9/PAS10) products, respectively. e  Amplicon sizes of 515/277 bp correspond to first round (Lip1/Lip4) and second round (Lip1/Lip2) products, respectively. The SNt-PCR detects both P. acnes and P. avidum. f  Second round amplicon size after Nt-PCR with PP-1/PP-2. First round PCR utilizes universal 16S rRNA primers to give a ~1500 bp product (see Section 2.2.2). g Amplicon sizes of 387/160 bp correspond to first round (Pa1/rPa3) and second round (Pa1/rPa2) products, respectively. The SNt-PCR will also give an amplicon with Actinomyces israelii, but the product can be differentiated from the P. acnes product by digestion with XhoI.

P. acnes

P. acnes

P. propionicum

P. acnes/ P. avidum

P. acnes

P. acnes

P. acnes

P. granulosum

P. acnes

P. granulosum

P. acnes

P. acnes

P. acnes

Species Targeted

TABLE 13.2 Species-Specific Oligonucleotide Primers and Probes for the Detection of Propionibacterium Species

144 Molecular Detection of Human Bacterial Pathogens

145

Propionibacterium

sequences.124 As mentioned earlier, nucleotide sequence analysis of the P. acnes recA and tly gene loci facilitates unambiguous determination of phylotype status (IA, IB, II, III).22,30 Specific PCR primers have now also been described for differentiation of P. acnes phylotypes, with type IA-specific primers targeting a solute-binding protein (SBP) gene, and type IB primers amplifying part of a putative Y40U fragment which brackets a point of sequence divergence between type IB and types IA and II. Type II can be differentiated from IA and IB by primers, which amplify within putative RHSfamily proteins, and bracket a ~3450-bp deletion in type II.66 Molecular methods have also been applied to propionibacteria in an attempt to differentiate the species and type individual strains. These methods have included ribotyping, Random Amplification of Polymorphic DNA (RAPD), which is a low stringency PCR that uses one or sometimes several primers of short length to generate a unique genetic fingerprint, and Amplified Fragment Length Polymorphism Analysis (AFLP), a technique based on the selective amplification of DNA fragments produced from a total restriction enzyme digest of genomic DNA.136–138 Macrorestriction digestion of genomic DNA followed by pulse field gel electrophoresis has also been used to type propionibacteria isolates.64,95,139







13.2  METHODS 13.2.1  Sample Preparation 13.2.1.1  P  reparation of Sonicate from Retrieved Medical Devices To significantly improve the detection of P. acnes infections associated with indwelling medical devices, such as prosthetic joints, catheters, and heart valves, it is imperative that samples are processed under strict anaerobic conditions and adherent bacteria removed by sonication. The procedure described is based on a modification of Tunney et al.140 Collection of Devices 1. When immediately removed from the patient, each component of an implant (e.g., femoral and acetabular components of a prosthetic hip) should be placed aseptically into a sterile bag and inserted into an anaerobic jar (e.g., Oxoid Anaerobic Jar HP0011, Oxoid Ltd.) containing a catalyst (e.g., Oxoid Low Temperature Catalyst BR0042, Oxoid Ltd.) and a Gaspak (e.g., Oxoid Gas Generating Kit BR0038, Oxoid Ltd.). 2. The jar should be transported immediately to the laboratory and placed inside an anaerobic cabinet (e.g., Don Whitley Mk III, Don Whitley Scientific Ltd.) containing an atmosphere of 80% N2, 10% H2, and 10% CO2 (v/v/v), where it can be opened. Sample Processing 1. Pre-reduce quarter-strength Ringer’s solution (25%, v/v) by the addition of l-cysteine (0.05%,  w/v) and



remove oxygen by boiling at 100°C for 1 h or micro­ waving at full power for 5 min. Upon cooling, store the solution in the anaerobic cabinet prior to use. 2. Add pre-reduced Ringer’s solution (100 mL) to the sterile bag containing the removed device and seal the bag with a sterile plastic clip. Gently agitate the bag by hand for 1–2 min to remove any extraneous blood and tissue and aspirate and discard the resulting solution. Add fresh sterile pre-reduced Ringer’s solution (100 mL) and reseal the bag. 3. Remove the bag from the cabinet and dislodge any adherent bacteria on the implant into the Ringer’s solution by mild ultrasonication (5 min in a 150 W ultrasonic bath operating at a frequency of 50 kHz). Transfer the Ringer’s solution from the sealed bag into 50 mL sterile centrifuge tubes inside the cabinet. Centrifuge the tubes for 20 min at 10,000 × g and 4°C and transfer back to the cabinet. Collectively resuspend the resulting pellets in a total volume of 5  mL Ringer’s solution (twenty fold concentration). 4. Total viable counts are performed by spread plating (within the anaerobic cabinet) 0.5 mL volumes of the resulting concentrated sonicate sample onto 3 × blood agar plates [BA; CM0271; Oxoid Ltd.; 5% (v/v) horse blood] and 3 × anaerobic blood agar plates [ABA; CM0972; Oxoid Ltd.; 5% (v/v) horse blood], which have been stored in the anaerobic cabinet prior to use. Incubate all BA and ABA plates at 37°C aerobically and anaerobically, respectively, for up to 14 days. The use of BA plates will allow the simultaneous identification of pathogenic aerobes in addition to propionibacteria and other anaerobes. 5. For molecular detection, extract DNA from 1 mL of sonicate using the methods described below. Store any remaining sample at –80°C for further analyses.

13.2.1.2  D  NA Extraction from Clinical and Bacterial Samples Different methods have been used for the extraction of bacterial DNA from clinical samples associated with propionibacteria infections, including prostate, lymph node, liver and prosthetic joint-associated tissue, vitreous and aqueous humors, root canal and sonicate samples, and bronchoalveolar lavage cells (Table 13.3).41,49,51,53,63,118–120,125–127,129,140–142 DNA has been extracted from freshly collected samples and has also been isolated from paraffin-embedded tissue samples. Methods have included the preparation of “crude” DNA from root canal samples by simple boiling to purification of genomic DNA from tissue; the latter often based on the use of commercially available kits. These methods can also be applied to the isolation of DNA from pure cultures of propionibacteria. (i) Purification of Bacterial Genomic DNA  The following method is based on that previously described by Tunney et al.

146

Molecular Detection of Human Bacterial Pathogens

TABLE 13.3 Summary of Different Methods Used to Extract DNA from Clinical Samples Associated with Propionibacterium Infections Referencea 53 118

Extraction Methodb

129 142 69,125 126

Qiagen QIAamp DNA mini Kit Roche MagNA Pure Compact Nucleic acid isolation kit Qiagen QIAamp DNA mini Kit; Phenol:Chloroform extraction Qiagen QIAamp DNA mini Kit; MasterPure DNA purification kit Qiagen QIAamp DNA mini Kit; Puregene DNA purification kit; Phenol:Chloroform extraction DEXPATTM kit and analysis of crude extracta Qiagen QIAamp DNA Blood Mini Kit Phenol:Chloroform extraction Boiling and analysis of a crude supernatant Qiagen QIAamp Tissue kit

41

Phenol:Chloroform extraction

50 120

Phenol:Chloroform extraction DEXPATTM kit and Phenol:Chloroform extractiona Phenol:Chloroform extraction

141 119

63

127 a b

Sample Types

Prosthetic joint infection Bacterial endophthalmitis; corneal ulcers Sarcoidosis

Core samples from thawed prostate tissue and seminal vesicle samples Section of human prostate tissue from frozen tissue block

Adenocarcinoma of the prostate gland

Laser capture microdissection of paraffin-embedded human liver tissue Section of paraffin-embedded human prostate tissue Human vitreous fluid Human root canal samples Section of paraffin-embedded human lymph node tissue Section of paraffin-embedded human lung and lymph node tissue Sonicate from failed prosthetic hip implant Section of paraffin-embedded human lymph node tissue Human vitreous fluid; aqueous humor

Primary biliary cirrhosis









Benign prostatic hyperplasia Uveitis associated with sarcoidosis Periodontitis Sarcoidosis Sarcoidosis Prosthetic joint infection Sarcoidosis Bacterial endophthalmitis

References are given in chronological order starting with most recent. DEXPATTM = DNA Extraction from Paraffin-embedded Tissue.

for the purification of bacterial DNA from sonicate samples but could be applied to other sample types or pure cultures.140

Disease

Sonicate from failed prosthetic shoulder implant Human vitreous fluid; aqueous humor; cell suspensions Bronchoalveolar lavage cells

1. Centrifuge 1 mL of sample at 10,000 × g for 15 min at room temperature and discard the supernatant. 2. Add lysozyme (1–2 mg/mL) in 10 mM Tris pH 8.0 and 50 mM glucose to aid extraction of the bacterial DNA. 3. Resuspend the pellet by vortex mixing in 200 μL cell lysis buffer [10 mM Tris pH 8.0, containing 5  mM EDTA, 0.5% (w/v) sodium dodecyl sulfate, and 100 μg/mL proteinase K] and incubate for 3 h at 55°C. Adjust the temperature to 37°C and leave overnight. The next morning, raise the temperature again to 55°C for 1 h. 4. Add an equal volume of saturated phenol/chloroform to the sample and mix by inversion before centrifugation at 10,000 × g for 15 min. This step can be repeated. 5. Remove the aqueous supernatant to a fresh tube making sure the interface is left behind. 6. Add ethanol (100%; 2.5 vols), 3 M sodium acetate, pH 5.2 (0.1 vol), and 2 μL SeeDNA (GE Healthcare) and mix by inversion followed by centrifugation at 10,000 × g for 15 min at room temperature. 7. Discard the supernatant and wash the pelleted DNA with 1 mL of 70% (v/v) ethanol. Centrifuge the



sample at 10,000 × g for 15 min at room temperature and remove the resulting supernatant. Dry the pellet in a vacuum dryer. 8. Dissolve the extracted DNA in TE buffer (10 mM Tris pH 8.0, 1 mM EDTA) and determine the concentration by UV absorbance at 260/280. The sample can be stored at –20°C.

Note: In previous studies with pure cultures, P. acnes has been grown in the presence of penicillin G to help weaken its thick gram-positive cell wall prior to DNA extraction.137 The use of additional enzymes such as lysostaphin or achromopeptidase in addition to lysozyme may improve cell wall lysis of propionibacteria.123 (ii) Preparation of Bacterial Genomic DNA by Boiling  If DNA from propionibacteria isolates is intended only for PCR, then preparation by boiling is much faster, unless a high-quality DNA template is required for PCR.66



1. Wash a small loop full of bacteria by suspension in 1 mL of phosphate buffered saline, pH 7.2, followed by centrifugation at 10,000 × g for 5 min to pellet the bacteria. 2. Resuspend the pellet in 100–200 μL of ultrapure water, boil for 10 min, and then chill on ice.

147

Propionibacterium



3. Centrifuge at 15,000 × g for 2 min to remove cell debris and store the supernatant at –20°C pending analysis.

13.2.1.3  P  reparation of Samples for Fluorescent In Situ Hybridization Preparations of samples for FISH are as described by Poppert et al. for pure bacterial cultures and by Alexeyev et al. for sections of tissue.117,123





1. For analysis of pure cultures, bacteria can be grown in broth or agar plates. Wash the bacteria and resuspend to the required concentration in 0.9% (w/v) saline. Apply approximately 10 μL of the bacterial suspension to an 8 well-glass slide, air dry, and fix for 10 min in pure methanol. Note: as a guide, an OD (600 nm) of 0.3 corresponds to approximately 108 CFU/ml for P. acnes. Remove the slides, air dry, and analyze immediately or store at –20°C for several weeks. 2. To help permeabilize the peptidoglycan layer prior to hybridization, treat fixed cultures with 6 μL of a solution of lysozyme (2 mg/mL) and lysostaphin (0.1 mg/ mL) in 10 mM Tris-HCl pH 8.0, for 10 min at 36°C. Stop the reaction by incubation in methanol for 3 min. 3. For tissue sections embedded in paraffin, dewaxing in xylene is mandatory. Dewax specimens in 100% xylene for 20 min and then rehydrate in an ethanol series (100%, 70%, 30%, and 0%) in 10 mM Tris, pH 7.5. Treat sections with lysozyme (1 mg/mL; Sigma-Aldrich) and achromopeptidase (30 U/mL; Sigma-Aldrich) in 10 mM Tris, pH 7.5 at 37°C for 25 min, wash for 5 min in 10 mM Tris, pH 7.5; and dehydrate in an ethanol series (30%, 70%, and 100%) in 10 mM Tris, pH 7.5.

13.2.2  Detection Procedures 13.2.2.1  PCR Detection of Propionibacterium Species Principle.  Based on the primers and protocols described for the detection of P. acnes and/or P. granulosum in human prostate tissue, sarcoid tissue and vitreous fluid from sarcoidic eyes with uveitis, and P. propionicum in human root canal samples.69,119,120,121,122 Procedure 1. The PCR mixture consists of 1× PCR buffer (20 mM Tris-HCl, pH 8.4, 50 mM KCl), 200 μM dNTPs, 300 nM each primer for P. acnes (PA-F’/PA-R) and P. granulosum (PG-1/PG-2) (Table 13.2) and 1 μM each primer for P. propionicum (PP-1/PP-2; Table 13.2), 2 mM MgCl2, 1.25 U Platinum Taq Hotstart DNA polymerase (Invitrogen Life Technologies), 1 μg of extracted genomic DNA and ultrapure water to a final volume of 25 μL. A negative control sample (water in place of DNA) should be included in each PCR run.





2. Run the following programs on a thermal cycler: for P. acnes and P. granulosum, initial denaturation at 95°C for 3 min; 40 cycles of 95°C for 60 s, primer pair annealing temperature (Table 13.2) for 60 s, 72°C for 60 s; and a final extension step at 72°C for 10 min. For P. propionicum, initial denaturation at 95°C for 2 min, 36 cycles of 95°C for 30 s, 55°C for 60 s, 72°C for 60 s; and a final extension step at 72°C for 2 min. 3. After PCR amplification, analyze 5–10 μL of each amplicon on a 1.5% (w/v) agarose gel in 1× Tris-acetate-EDTA buffer containing 0.5 μg/mL ethidium bromide or, alternatively, 1:10,000 dilution of GelRed (Cambridge Bioscience). A DNA molecular size marker should be run alongside and the resolved products visualized and photographed under UV light.

Note: PCR method can be applied to clinical samples and used to identify isolates. 13.2.2.2  N  ested and Semi-Nested PCR Detection of Propionibacterium propionicum and Propionibacterium acnes Principle.  Based on the primers and protocols described for nested-PCR detection of Propionibacterium propionicum in human root canal samples and semi-nested PCR detection of Propionibacterium acnes in human vitreous samples.125,128 Procedure 1. For nested-PCR detection of P. propionicum, the first round consists of the amplification of nearly full length 16S rRNA using the universal forward primer 5′-AGAGTTTGATCCTGGCTCAG-3′ and reverse primer 5′-ACGGCTACCTTGTTACGACTT-3′. The PCR mixture consists of 1× PCR buffer (20 mM Tris-HCl pH 8.4, 50 mM KCl), 200 μM dNTPs, 200 nM each primer, 2 mM MgCl2, 1.25 U Tth DNA polymerase (Biotools), 2 μL DNA preparation, and ultrapure water to a final volume of 25 μL. A negative control sample (water in place of DNA) should be included in each PCR run. 2. Run the following program on a thermal cycler: initial denaturation at 97°C for 60 s; 26 cycles of 97°C for 45 s, 55°C for 45 s, 72°C for 60 s; and a final extension step at 72°C for 4 min. 3. After first round amplification, 1 μL of the resulting product is used as the template for the nested species-specific reaction. PCR set up conditions are as described in (1) but with 100 nM of the P. propionicum primers PP-1/PP-2 (Table 13.2). Amplification conditions are initial denaturation at 95°C for 2 min; 26 cycles of 95°C for 30 s, 55°C for 60 s, 72°C for 60 s; and a final extension step at 72°C for 2 min. 4. For seminested PCR detection of P. acnes, the first round PCR mixture consists of 1×  PCR buffer (10 mM Tris-HCl, pH 8.3, 50 mM KCl, 0.01% (w/v) gelatin, 1.5 mM MgCl2), 200 μM dNTPs, 1 μM each of primers Pa1/rPa3 (Table 13.3), 0.5 U Taq DNA polymerase

148







(Stratagene), 10 ng DNA, and ultrapure water to a final volume of 25 μL. A negative control sample (water in place of DNA) should be included in each PCR run. 5. Run the following program on a thermal cycler: initial denaturation at 94°C for 5 min; 34 cycles of 94°C for 30 s, 62°C for 60 s, 72°C for 2 min; and a final extension step at 72°C for 8 min. 6. After first round amplification, 1 μL of the resulting product is used as the template for the seminested reaction. PCR set up and cycling conditions are as described in (4) and (5) but using the primer combination Pa1/rPa2 (Table 13.2) and only 24 amplification cycles. 7. After PCR amplification, analyze 5–10 μL of each amplicon on a 1.5% (w/v) agarose gel in 1 × Trisacetate-EDTA buffer containing 0.5 μg/mL ethidium bromide or, alternatively, 1:10,000 dilution of GelRed (Cambridge Bioscience). A DNA molecular size marker should be run alongside and the resolved products visualized and photographed under UV light.

Note: When setting up the PCR master mix for broad-range 16S rRNA PCR, all reagents except the primers and template DNA can be UV irradiated for 15 min to destroy any artificial DNA.143 This may, however, reduce the detection limit of the PCR. Particular care must also be taken to ensure that samples are not contaminated by “carry over” products from first round amplification. The seminested PCR described for P. acnes will also generate an amplicon from Actinomyces israelii, but the product can be differentiated by digestion with the restriction enzyme XhoI. 13.2.2.3  F luorescent In Situ Hybridization for the Detection of Propionibacterium acnes Principle.  Based on the hybridization protocol described for the detection of P. acnes in prostate tissue.123 Procedure 1. Incubate approximately 100–120 ng each of the P.  acnes-specific Cy3.5-tagged probe PAC23S (Table 13.2), plus four additional P. acnes-specific probes generated by PCR amplification and cloning (see notes), all in 10 µL of 40 mM sodium phosphate buffer, pH 7.0, containing 50% (v/v) formamide, 2× Saline-Sodium Citrate buffer (SSC) (1× SSC is 0.15 M NaCl plus 0.015 M sodium citrate), 10% (w/v) dextran sulfate, 0.1% (w/v) sodium dodecyl sulfate, and 1× Denhardt’s solution, on the slide at 75°C for 5 min, followed by overnight incubation at 37°C. Denhardt’s solution comprises high molecular weight polymers that are capable of saturating nonspecific binding sites and artificially increasing the concentration of probe available for hybridization. 2. After hybridization, wash the slides with 2% SSC containing 0.3% Nonidet P-40 (Biochem) and stain with 4,6-diamidino-2-phenylindole dihydrochloride (DAPI; Sigma).

Molecular Detection of Human Bacterial Pathogens





3. Wash off the DAPI stain from the slide with phosphate buffered saline and allow to air dry. Mount the slides in Citi-fluor antifade solution (Citi-fluor, England) and seal coverslips with nail varnish. 4. Examine slides under a fluorescence microscope and capture different fields of view electronically.

Note: In addition, Alexeyev et al. also utilized four large P. acnes probes (size range, 5–6 kb) that overlapped a 14,313 bp segment of the KPA171202 genome.123 These four fragments were synthesized by using Elongase enzyme mix (Invitrogen) and the following primer sets: Primers 1L (CCAAAGACAATTCTGGGAAGATTACAG, nucleotides 320829–320855) and 1R (ATACAACTTC GGCCTTTACACTACAGC, nucleotides 326085–326059); Primers 2L (AAAATGTGAAGTGTGAGGTGATTCTGC, nucleotides 325533–325559) and 2R (ATTAAGGTG GAATGGTTCTACTGGTTG, nucleotides 331266–331240); Primers 3L (ATAATGAGGATGGTGCAGACCAGTAG, nucleotides 330317–330342) and 3R (GATGGAGGA AAT CACATATAGCGAGAC, nucleotides 335700–335674); Primers 4L (GAGTGGATCTTCTTCGTCAACGTC, nucleotides 335119–335142) and 4R (AGAACTCGTCCTTCTTCA ACTTCCAAC; nucleotides 341090–341064). Products were cloned by using a Topo XL PCR cloning kit (Invitrogen) and the plasmids isolated by use of the QIAGEN midi kit. The libraries were then directly labeled with diethylaminocoumarin-dUTP (DEC) (Perkin-Elmer) by using a nick translation kit (Vysis Inc.). 13.2.2.4  N  ucleotide Sequence Analysis for Differentiation of Propionibacterium acnes Phylotypes Principle.  Based on sequence analysis of the recA housekeeping gene and the putative hemolysin/cytotoxin gene tly.22,30 Procedure 1. The PCR mixture consists of 1× PCR buffer (20 mM Tris-HCl pH 8.4, 50 mM KCl), 200 μM dNTPs, 200 nM each of primers PAR-1/PAR-2 (Table 13.4), 1.5 mM MgCl2, 1.25 U Platinum Taq Hotstart DNA polymerase (Invitrogen), 2 μL of DNA (prepared by boiling method), and ultrapure water to a final volume of 25 μL. A negative control sample (water in place of DNA) should be included in each PCR run. 2. Run the following program on a thermal cycler: one cycle at 95°C for 3 min; 35 cycles of 95°C for 60 s, 55°C for 30 s, 72°C for 90 s; and a final extension step at 72°C for 10 min. 3. After PCR amplification, analyze 5–10 μL of each amplicon on a 1.5% (w/v) agarose gel in 1× Tris-acetate-EDTA buffer containing 0.5 μg/mL ethidium bromide or, alternatively, 1:10,000 dilution of GelRed (Cambridge Bioscience). A DNA molecular size marker should be run alongside and the resolved products visualized and photographed under UV light.

149

Propionibacterium







4. Prior to sequencing, purify PCR products (5–10 μL) using a QIA-quick PCR purification kit (Qiagen). This kit provides spin columns, buffers, and collection tubes for silica membrane-based purification of PCR products >100 bp in size. After purification, estimate the concentration of the PCR product by running a small amount (1 μL) on a 1.5% (w/v) agarose gel (as described earlier) alongside a DNA ladder with known mass amounts per fragment. 5. To sequence, use a BigDye Terminator v 3.1 Cycle Sequencing Kit, which provides all the necessary reagents in a premixed, ready reaction format (Applied Biosystems). As described by the manufacturer, the sequencing mixture should consist of ×1 BigDye sequencing buffer, ×1 Ready Reaction Premix, 3.2 pmol of the relevant sequencing primer, 10–20 ng of the purified PCR product and ultrapure water to a final volume of 20 μL, all in a 0.2 mL microcentrifuge tube. Keep all samples on ice during preparation. Place samples on a thermocycler and carry out sequencing under the following conditions: one cycle at 96°C for 1 min; 25 cycles of 96°C for 10 s, 50°C for 5 s, 60°C for 4 min. 6. Purify sequenced samples using a Montage SEQ96 Sequencing Reaction Cleanup Kit (Millipore) according to the manufacturer’s instructions and analyze on an ABI PRISM® analyzer, such as the ABI PRISM® 3100 genetic analyzer. After analysis, open the electropherogram files using a sequence analysis software program such as Chromas or EditView. Phylotype status can be determined by comparison of resulting sequences with those in online databases.

Note: The primers PAR-1 and PAR-2 target sequences which flank the 5′ and 3′ ends of the recA open reading frame, respectively, while PAT-1 and PAT-2 similarly target sequences that flank the 5′ and 3′ ends of the tly open reading frame. Complete gene sequences can be obtained by using both the forward and reverse primers for each locus. Typespecific polymorphisms in the recA and tlyA genes will facilitate phylotyping of each isolate; type I isolates (IA and IB)

differ from type II in 10 specific bases within the recA locus, while the tly gene shows greater variation with 18 specific differences between type I (IA and IB) and type II.30 Type IB strains contain an additional polymorphism at nucleotide position 897 in recA and two polymorphisms at positions 375 and 766 in tly (which match those in type IIs) compared to type IA. Some strains of IB may also contain an additional polymorphism in recA (McDowell & Patrick, unpublished). Strains within the type III clade contain a combination of four type I-specific polymorphisms (common to IA and IB), six type II-specific polymorphisms, and five polymorphisms unique to the group.22 13.2.2.5  P  CR Differentiation of Propionibacterium acnes Types IA, IB, and II Principle.  Based on the method of Shannon et al.66 Variations in DNA sequence between types IA, IB, and II were detected by RAPD using a range of random primers. Amplified bands specific to each phylotype fingerprint were then sequenced and used as the basis for the design of typespecific primers. Procedure 1. The PCR mixture consists of 1× PCR buffer containing 1.5 mM MgCl2 (Qiagen), 200 μM dNTPs, 500 nM each primer (Table 13.4), 0.75 U Hotstar Taq DNA polymerase (Qiagen), 2 μL of purified genomic DNA (Qiagen DNA Extraction Kit), and ultrapure water to a final volume of 20 µL. A negative control sample (water in place of DNA) should be included in each PCR run. 2. Run the following program on a thermal cycler: 15 min at 95°C to activate the HotStar Taq; 35 cycles of 30 s at 94°C, 30 s at annealing temperature (65°C for PR245/247; 55°C for the other primer sets), 72°C for 1 min; and a final extension step at 72°C for 10 min. 3. After PCR amplification, analyze 10–20 μL on a 1.5% (w/v) agarose gel in 1× Tris-acetate-EDTA buffer containing 0.5 mg/mL ethidium bromide or, alternatively, 1:10,000 dilution of GelRed (Cambridge

TABLE 13.4 PCR Primers for Phylotyping of Propionibacterium acnes Isolates Reference 22,30

Specificity

Primer

Sequences (5′–3′)

All phylotypes

PAR-1 PAR-2 PAT-1 PAT-2 PR213 PR216 PR256 PR257 PR245 PR247

AGCTCGGTGGGGTTCTCTCATC GCTTCCTCATACCACTGGTCATC CAGGACGTGATGGCAATGCGA TCGTTCACAAGACCACAGTAGC GACGATTACTGGGGCAAGAA GGGGTGAACTTGACGTTGAT GTCATCGGCTTCTGGATCAT CCTGGTTTCCACGATGTCTT ATAAACCCATCGGCGGCTCGAT AGCAAGGGAGCATTCAGTGG

All phylotypes 66

Type IA Type IB Type IA, IB, II

Amplicon Size (bp)

Target Loci

Note

1201

recA

909

tly

584 725 4027 (IA, IB) 573 (II)

sbp Y40U RHS

Identify phylotype by nucleotide sequencing of amplicons. Primers target sequences flanking the open reading frames of both genes. With the primer pair PR245/247, type II isolates generate a much smaller PCR product compared with IA and IB

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Molecular Detection of Human Bacterial Pathogens

Bioscience). A DNA molecular size marker should be run alongside and the resolved products visualized and photographed under UV light. Note: For optimal performance of these type-specific PCRs, and especially for successful amplification of the 4027-bp fragment from types IA and IB with PR245/247, it is important that purified genomic DNA is used as a good quality template. The primers PR213/PR216 amplify a portion of the solute binding-protein gene (sbp) of a peptide uptake operon that is within an 8.7-kb region present between nucleotides 1579146 and 1579405 in type IA isolates only (GenBank accession number DQ208967). The primer combination of PR256/PR257 amplify part of a putative Y40U fragment (nucleotides 948634–949358), which brackets a region of sequence divergence between types IA and IB, with the PR256 binding site being present only in type IB and either absent or significantly different in sequence in type II. Finally, the primer pair PR245/PR247 amplify putative RHS-family proteins (nucleotide 246705–250731) and bracket a ~3450-bp deletion in type II, giving rise to a smaller amplicon compared to types IA and IB.

13.3  C  ONCLUSIONS AND FUTURE PRESPECTIVES Within the genus Propionibacterium, only those organisms that are members of the “cutaneous” group are associated with human disease. Although P. granulosum and P. avidum have been found to cause opportunistic human infections, in practical terms it is P. acnes and to a lesser extent P. propionicum that are associated with the overwhelming majority of propionibacteria infections in humans. The relationship between the “cutaneous” propionibacteria and humans is, under normal circumstances, a positive one. They are an important component of the normal microbiota on human skin where they help maintain a healthy ecosystem and prevent colonization with more pathogenic microbes.144 Due to their potent immunomodulatory properties, their use as adjuvants may also be beneficial in the treatment of various cancers.145 In terms of disease, P. acnes is probably the most well-known species within the Propionibacterium genus due to its association with acne. There is, however, a growing body of evidence to suggest that its involvement in other infections and diseases has been significantly underestimated, especially in relation to infections of indwelling medical devices where it will persist as an adherent biofilm.49,53 This has occurred, in part, due to the traditional and overly simplistic view that P. acnes is a relatively harmless member of the human microbiota and therefore its presence in clinical samples simply reflects contamination. In addition, failure of prosthetic orthopaedic implants due to infection is often not suspected with P. acnes (or looked for postoperatively) due to subtle signs and symptoms and, consequently, patients are often misdiagnosed preoperatively as presenting with aseptic loosening. To compound this, in some laboratories anaerobic processing and culture of

specimens may either not occur or be inappropriate resulting in poor diagnostic sensitivity for P. acnes and other propionibacterial infections. To improve detection, samples should be processed under strict anaerobic conditions, retrieved indwelling devices sonicated to ensure adherent organisms are removed, and cultures incubated for at least 7 days, and ideally up to 14 days, to prevent a false-negative result. While culture-based detection remains the mainstay of the diagnostic bacteriologist, molecular-based analysis of specimens is a valuable adjunct and, due to much shorter processing times, can lead to faster diagnosis as well as the capacity to identify VBNC organisms. This may be especially important in relation to prosthetic hip joint revision arthroplasty, where rapid and reliable identification of propionibacteria infection on the removed device may significantly impact on patient treatment and management. If available, the application of real-time PCR for the detection of propionibacteria in samples can offer notable improvements on conventional PCR methods due to reduced assay times, postamplification analysis, and increased specificity and sensitivity. Background amplification products can also be identified based on quantification and melting curve analysis. Problems can be encountered when attempting to optimally extract DNA from propionibacteria due to the thickness of their gram-positive cell envelope and low sensitivity to lysozyme and proteinase K. This could impact on detection limits for PCR or FISH diagnosis when using tissue samples and therefore attention should be paid to the use of appropriate enzymatic treatments that may be more appropriate for lysis, such as achromopeptidase.123 The molecular identification of novel phylotypes within P. acnes has stimulated investigations of whether these distinct groups vary in their ability to cause disease, and are, therefore, clinically important. Their discovery has also raised important issues regarding how we now compare and contrast data generated in previous P. acnes-related studies where the phylotype status of isolates was not known. Continued and improved molecular epidemiological studies will hopefully help to address these issues. The molecular detection of phylotypes associated with greater pathogenicity could prove especially valuable information for the diagnostic bacteriologist as they attempt to confirm whether a P. acnes isolate is clinically relevant. The availability of the P. acnes complete genome sequence will undoubtedly facilitate the development of improved molecular detection and typing platforms for the bacterium, and it is hoped that the complete genome sequences of other “cutaneous” species and P. propionicum will eventually be determined. This will facilitate direct genomic comparisons, give important and much needed insights into the pathogenic potential of these other organisms, and provide a foundation for improved diagnostics. The development of DNA-microarrays based on such Propionibacterium genome sequences would also facilitate transcriptomic profiling and the potential identification of propionibacterial genes upregulated during disease. These loci could then provide novel diagnostic platforms for the detection of infection versus contamination by reverse-transcriptase PCR.

Propionibacterium

ACKNOWLEDGMENTS The authors acknowledge the Northern Ireland Health and Social Care Research & Development Office (NIHSC R&D Office), Arthritis Research UK, the British Orthopaedic Society Wishbone Trust and The Prostate Cancer Charity UK, for funding to investigate the role of Propionibacterium acnes in orthopaedic-related infections and prostate cancer.

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Propionibacterium 77. Werner, H., Lipase and lecithinase activities of aerobic and anaerobic Corynebacterium and Propionibacterium species, Zentralbl. Bakteriol. Orig., 204, 127, 1967. 78. Smith, R.F., and Willett, N.P., Lipolytic activity of human cutaneous bacteria, J. Gen. Microbiol., 52, 441, 1968. 79. Ingham, E. et al., Partial purification and characterization of lipase (EC 3.1.1.3) from Propionibacterium acnes, J. Gen. Microbiol., 124, 393, 1981. 80. Ingram, E. et al., Studies of the extracellular proteolytic activity produced by Propionibacterium acnes, J. Appl. Bacteriol., 54, 263, 1983. 81. Brüggemann, H. et al., The complete genome sequence of Propionibacterium acnes, a commensal of human skin, Science, 305, 671, 2004. 82. Valanne, S. et al., CAMP factor homologues in Propionibacterium acnes: A new protein family differentially expressed by types I and II, Microbiology, 151, 1369, 2005. 83. Lang, S., and Palmer, M., Characterization of Streptococcus agalactiae CAMP factor as a pore-forming toxin, J. Biol. Chem., 278, 38167, 2003. 84. Brüggemann, H., Insights in the pathogenic potential of Propionibacterium acnes from its complete genome, Semin. Cutan. Med. Surg., 24, 67, 2005. 85. Lodes, M.J. et al., Variable expression of immunoreactive surface proteins of Propionibacterium acnes, Microbiology, 152, 3667, 2006. 86. Hammerschmidt, S. et al., Capsule phase variation in Neisseria meningitidis serogroup B by slipped-strand mispairing in the polysialyltransferase gene (siaD): Correlation with bacterial invasion and the outbreak of meningococcal disease, Mol. Microbiol., 20, 1211, 1996. 87. Fink, H. et al., Systemic inflammatory response syndrome increases immobility-induced neuromuscular weakness, Crit. Care Med., 36, 910, 2008. 88. Flaminio, M.J., Rush, B.R., and Shuman, W., Immunologic function in horses after non-specific immunostimulant administration, Vet. Immunol. Immunopathol., 63, 303, 1998. 89. Shannon, B.A., Garrett, K.L., and Cohen, R.J., Links between Propionibacterium acnes and prostate cancer, Future Oncol., 2, 225, 2006. 90. Graham, G.M. et al., Proinflammatory cytokine production by human keratinocytes stimulated with Propionibacterium acnes and P. acnes GroEL, Br. J. Dermatol., 150, 421, 2004. 91. Westerhof, W. et al., Propionibacterium acnes and the pathogenesis of progressive macular hypomelanosis, Arch. Dermatol., 140, 210, 2004. 92. Nagy, I. et al., Propionibacterium acnes and lipopolysaccharide induce the expression of antimicrobial peptides and proinflammatory cytokines/chemokines in human sebocytes, Microbes Infect., 8, 2195, 2006. 93. Tunney, M.M. et al., Biofilm formation by bacteria isolated from retrieved failed prosthetic hip implants in an in vitro model of hip arthroplasty antibiotic prophylaxis, J.  Orthop. Res., 25, 2, 2007. 94. Coenye, T., Peeters, E., and Nelis, H.J., Biofilm formation by Propionibacterium acnes is associated with increased resistance to antimicrobial agents and increased production of putative virulence factors, Res. Microbiol., 158, 386, 2007. 95. Oprica, C., and Nord, C.E., European surveillance study on the antibiotic susceptibility of Propionibacterium acnes, Clin. Microbiol. Infect., 11, 204, 2005. 96. Ross, J.I. et al., Antibiotic-resistant acne: Lessons from Europe, Br. J. Dermatol., 148, 467, 2003.

153 97. Tan, H.H. et al., Antibiotic sensitivity of Propionibacterium acnes isolates from patients with acne vulgaris in a tertiary dermatological referral centre in Singapore, Ann. Acad. Med. Singapore, 30, 22, 2001. 98. Ishida, N. et al., Antimicrobial susceptibilities of Propionibacterium acnes isolated from patients with acne vulgaris, Microbiol. Immunol., 52, 621, 2008. 99. Chow, A.W., Patten, V., and Guze, L.B., Susceptibility of anaerobic bacteria to metronidazole: Relative resistance of non-spore-forming gram-positive baccilli, J. Infect. Dis., 131, 182, 1975. 100. Pochi, P.E., and Strauss, J.S., Antibiotic sensitivity of Corynebacterium acnes (Propionibacterium acnes). J. Invest. Dermatol., 36, 423, 1961. 101. Wang, W.L. et al., Susceptibility of Propionibacterium acnes to seventeen antibiotics, Antimicrob. Agents Chemother., 11, 171, 1977. 102. Hoeffler, U., Ko, H.L., and Pulverer, G., Antimicrobiol susceptibility of Propionibacterium acnes and related microbial species, Antimicrob. Agents Chemother., 10, 387, 1976. 103. Eady, E.A. et al., Erythromycin resistant propionibacteria in antibiotic treated acne patients: Association with therapeutic failure, Br. J. Dermatol., 121, 51, 1989. 104. Malik, A.C., Reinbold, G.W., and Vedamuthu, E.R., An evaluation of the taxonomy of Propionibacterium, Can. J. Microbiol., 14, 1185, 1968. 105. Kishishita, M. et al., New medium for isolating propionibacteria and its application to assay of normal flora of human facial skin, Appl. Environ. Microbiol., 40, 1100, 1980. 106. Jousimies-Somer, H. et al., Wadsworth-KTL Anaerobic Bacteriology Manual, 6th ed., Star Publishing, Belmont, 2002. 107. Bjerkan, G., Witsø, E., and Bergh K., Sonication is superior to scraping for retrieval of bacteria in biofilm on titanium and steel surfaces in vitro, Acta Orthop., 80, 245, 2009. 108. Mordarska, H., and Pas´ciak, M., A simple method for differentiation of Propionibacterium acnes and Propionibacterium propionicum, FEMS. Microbiol. Lett., 123, 325, 1994. 109. Celig, D.M., and Schreckenberger, P.C., Clinical evaluation of the RapID-ANA II panel for identification of anaerobic bacteria, J. Clin. Microbiol., 29, 457, 1991. 110. Looney, W.J., Gallusser, A.J., and Modde H.K., Evaluation of the ATB 32 A system for identification of anaerobic bacteria isolated from clinical specimens, J. Clin. Microbiol., 28, 1519, 1990. 111. Cavallaro, J.J., Wiggs, L.S., and Miller, J.M., Evaluation of the BBL Crystal Anaerobe identification system, J. Clin. Microbiol., 35, 3186, 1997. 112. Funke, G. et al., Multicenter evaluation of the updated and extended API (RAPID) Coryne database 2.0, J. Clin. Microbiol., 35, 3122, 1997. 113. Oliver, J.D., The viable but nonculturable state in bacteria, J. Microbiol., 43, 93, 2005. 114. Brown, M.R., and Williams P., The influence of environment on envelope properties affecting survival of bacteria in infections, Annu. Rev. Microbiol., 39, 527, 1985. 115. Palomino, J.C. et al., Rapid culture-based methods for drug-resistance detection in Mycobacterium tuberculosis, J. Microbiol. Methods, 75, 161, 2008. 116. Janda, J.M., and Abbott, S.L., Bacterial identification for publication: When is enough? J. Clin. Microbiol., 40, 1887, 2002.

154 117. Poppert, S., Riecker, M., and Essig A., Rapid identification of Propionibacterium acnes from blood cultures by fluorescence in situ hybridization, Diagn. Microbiol. Infect. Dis., 66, 214, 2010. 118. Goldschmidt, P. et al., New test for the diagnosis of bacterial endophtalmitis, Br. J. Ophthalmol., 93, 1089, 2009. 119. Sfanos, K.S. et al., A molecular analysis of prokaryotic and viral DNA sequences in prostate tissue from patients with prostate cancer indicates the presence of multiple and diverse microorganisms, Prostate, 68, 306, 2008. 120. Ishige I. et al., Quantitative PCR of mycobacterial and propionibacterial DNA in lymph nodes of Japanese patients with sarcoidosis, Lancet, 354, 120, 1999. 121. Yasuhara, T., et al., The presence of Propionibacterium spp. in the vitreous fluid of uveitis patients with sarcoidosis, Acta. Ophthalmol. Scand., 83, 364, 2005. 122. Sfanos, K.S., and Isaacs, W.B., An evaluation of PCR primer sets used for detection of Propionibacterium acnes in prostate tissue samples, Prostate, 68, 1492, 2008. 123. Alexeyev, O.A. et al., Direct visualization of Propionibacterium acnes in prostate tissue by multicolor fluorescent in situ hybridization assay, J. Clin. Microbiol., 45, 3721, 2007. 124. Nakamura, M. et al., Identification of Propionibacterium acnes by polymerase chain reaction for amplification of 16S ribosomal RNA and lipase genes, Anaerobe, 9, 5, 2003. 125. Siqueira, J.F., Jr., and Rôças, I.N., Polymerase chain reaction detection of Propionibacterium propionicus and Actinomyces radicidentis in primary and persistent endodontic infections, Oral Surg. Oral Med. Oral Pathol. Oral Radiol. Endod., 96, 215, 2003. 126. Yamada, T. et al., In situ localization of Propionibacterium acnes DNA in lymph nodes from sarcoidosis patients by signal amplification with catalysed reporter deposition, J. Pathol., 198, 541, 2002. 127. Therese, K.L., Anand, A.R., and Madhavan, H.N., Polymerase chain reaction in the diagnosis of bacterial endophthalmitis, Br. J. Ophthalmol., 82, 1078, 1998. 128. Hykin, P.G. et al., The diagnosis of delayed post-operative endophthalmitis by polymerase chain reaction of bacterial DNA in vitreous samples, J. Med. Microbiol., 40, 408, 1994. 129. Alexeyev, O. et al., Association between the presence of bacterial 16S RNA in prostate specimens taken during transurethral resection of prostate and subsequent risk of prostate cancer (Sweden), Cancer Cause Control, 17, 1127, 2006. 130. Clarridge, III, J.E., Impact of 16S rRNA gene sequence analysis for identification of bacteria on clinical microbiology and infectious diseases, Clin. Microbiol. Rev., 17, 840, 2004.

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14 Rhodococcus Debarati Paul, Dipaloke Mukherjee, Soma Mukherjee, and Debosmita Paul CONTENTS 14.1 Introduction...................................................................................................................................................................... 155 14.1.1 Classification and Epidemiology.......................................................................................................................... 155 14.1.2 Clinical Features................................................................................................................................................... 156 14.1.3 Pathogenesis and Virulence.................................................................................................................................. 158 14.1.4 Cell Biology.......................................................................................................................................................... 160 14.1.5 Diagnosis.............................................................................................................................................................. 160 14.1.5.1 Phenotypic Identification....................................................................................................................... 160 14.1.5.2 Molecular Identification......................................................................................................................... 162 14.1.6 Treatment and Control.......................................................................................................................................... 162 14.2 Methods............................................................................................................................................................................ 164 14.2.1 Sample Preparation............................................................................................................................................... 164 14.2.2 Detection Procedures............................................................................................................................................ 164 14.3 Conclusion and Future Perspectives................................................................................................................................. 165 Acknowledgments...................................................................................................................................................................... 165 References.................................................................................................................................................................................. 165

14.1  INTRODUCTION The genus name “Rhodococcus” was first used by Zopf in 1891 to describe two species of red-pigmented bacteria.1–3 The taxon Rhodococcus is of interest for a variety of reasons, including its several metabolic abilities, as they can degrade a wide range of environmental pollutants and transform or synthesize a number of compounds with possible useful applications. Rhodococci are virtually ubiquitous, which is evident by their presence in a large variety of sources including soils, rocks, boreholes, groundwater, marine sediments, animal dung, the guts of insects, and in healthy and diseased animals and plants.4–6 The commercial potential of Rhodococcus species is increasingly being recognized. Their outstanding ability to synthesize several products such as surfactants, flocculants, amides, and polymers, as well as their capacity to degrade or transform a wide range of chemicals make rhodococci actually or potentially useful in environmental and industrial biotechnology. This increasing interest is reflected in patenting. Finnerty lists 10 patents relating to rhodococci up to 1990,1 Warhurst and Fewson report that a further 20 patent families were submitted in the following two years, while from 1993 to November 1996, a further 80 patents were submitted to the World Patent Index.6 Interestingly, two strains of Rhodococcus triatome have been identified as symbionts in bloodsucking bugs of the genus Triatoma.7 There has also been an increased interest in infections caused by rhodococci in humans, animals, and plants.4,8–10 Rhodococci are nonpathogenic with the exception

of two strains, namely R. fascians11 and R. equi.12 Rhodococcus fascians is the only described phytopathogen in this genus that causes leafy gall disease and is a biotrophic organism with broad host range. Through the secretion of potent morphogens, the bacteria stimulate the formation of new shoots from differentiated tissues.13–18 However, since it is out of the scope of this chapter, we would limit our discussion on this pathogen. The other pathogenic Rhodococcus is R. equi, which causes infections in both horses and humans. It is a dangerous pathogen of young foals aged up to 3–5 months.19

14.1.1  Classification and Epidemiology The current systematics of this genus in the NCBI database is as follows: Actinobacteria, Actinobacteridae, Actinomycetales, Corynebacterineae, Nocardiaceae, and Rhodococcus. However, this is unclear, partly because many members were originally included before the application of a polyphasic taxonomic approach, central to which is the acquisition of 16S rRNA sequence data. Bacteria of the order Actinomycetales were originally classified as fungi because of their hyphal and mycelial morphology, and include the mycolic acid-containing Corynebacterineae, which presently contain the genera Gordonia, Rhodococcus, and Nocardia.2 Rhodococcus was revived and redefined in 1977 to accommodate the “rhodochrous” complex, which comprised a number of strains that resembled but did not belong to the established genera Nocardia, Corynebacterium, and Mycobacterium. Rhodococci are described as aerobic, gram-positive, nonmotile 155

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(with exception of R. agilis), mycolate-containing, nocardioform actinomycetes. The term “nocardioform” is morphologically descriptive and refers to mycelial growth with fragmentation into rod-shaped or coccoid elements. There are currently 12 established Rhodococcus species, namely R. coprophilus, R. equi, R. fascians, R. erythropolis, R. globerulus, R. marinonascens, R. opacus, R. percolatus, R. rhodnii, R. rhodochrous, R. ruber, and R. zopfii.20 Some strains bearing other generic and specific names, such as Arthrobacter picolinophilus, Corynebacterium hoagii, Nocardia restricta, and Nocardia calcarea have been shown recently to belong to established Rhodococcus species. Nocardia corynebacteroides also appears to be a Rhodococcus but does not belong to any of the currently valid species.9,21,22 One of the most important characteristic features that mark the differences between Nocardia species from that of Rhodococcus is the presence of an MK-8(H4) type cyclic component in menaquinone. All the species under the genus Nocardia exhibits this feature with the exception of the strain N. corynebacteroides.9 Other invalidated species names, such as R. longus, R. minimus, and R. luganensis,2 occur in the literature, while there are many strains that are not identified to species level.2 In order to provide a framework for a taxonomic approach based on multiple genetic loci, a survey of the known genome characteristics of members of the genus Rhodococcus was undertaken. This included (i) genetics of cell envelope biosynthesis; (ii) virulence genes; (iii) gene clusters involved in metabolic degradation and industrially relevant pathways; (iv) genetic analysis tools; (v) rapid identification of bacteria including rhodococci with specific gene RFLPs; and (vi) genomic organization of rrn operons. Genes encoding virulence factors have been characterized for Rhodococcus equi and Rhodococcus fascians. Based on peptide signature comparisons deduced from gene sequences for cytochrome P-450, mono- and dioxygenases, alkane degradation, nitrile metabolism, proteasomes, and desulfurization, phylogenetic relationships can be deduced for Rhodococcus erythropolis, Rhodococcus globerulus, Rhodococcus ruber, and a number of undesignated Rhodococcus spp. that may distinguish the genus Rhodococcus into two further genera.3 R. equi is a gram-positive, pleomorphic, intracellular bacillus that is an obligatory aerobe, sometimes weakly acidalcohol resistant, and with distinctive biochemical features. R. equi is ubiquitous in soil, as well as in the feces of herbivores, birds, dogs, and so forth. It is a common pathogen in domestic animals that occasionally affects humans. The pathogenicity of R. equi is expressed mostly in hosts who are immunosuppressed.23 Infection in other animal species is rare and usually the result of immunosuppression. Rhodococcus equi is an important cause of AIDS-associated pneumonia in HIV-infected humans.24 This pathogen was originally isolated by Magnusson in 1923 from granulomatous lung infections in young horses.25 Infection in humans may result from environmental exposure26,27 because the organism is abundantly present in soil.28 The stool of horses and other animals is the source of soil contamination. While early cases occurred mostly in persons with a history of contact with

Molecular Detection of Human Bacterial Pathogens

horses, only 20%–30% of recent cases can be traced to such contact.29 R. equi infection in a human was first reported in 1967 in a 29-year-old man with plasma cell hepatitis who was receiving immunosuppressant medications. Since then, R. equi has become an important opportunistic pathogen in immunocompromised patients, especially those with AIDS.

14.1.2  Clinical Features R. equi is a rare opportunistic pathogen found in severely immunocompromised patients and also in human immunodeficiency virus (HIV)-infected persons, and is associated with significant mortality. In earlier cases, patients receiving immunosuppressant therapy were more likely to be successfully treated with antimicrobial agents as compared to the cases in AIDS patients.30 Most often, patients have a slowly progressive granulomatous pneumonia, with lobar infiltrates, frequently developing to cavitating lesions visible on chest X-ray (Figure 14.1a and 14.1b). Other sites of infection include abscesses of the central nervous system, pelvis, subcutaneous tissue, and lymphadenitis.29–31 Delays in accurate diagnosis of R. equi are still very common27,29 despite increased awareness of this organism as an opportunistic pathogen in humans. Factors for delayed diagnosis include the insidious onset of disease, clinical resemblance of the infection to mycobacterial, fungal, and actinomycotic infections, and the relatively nondescript bacteriologic profile of R. equi. Table 14.1 lists some of the previously reported clinical manifestations of R. equi infection. The sites of infection and the specimens from which R. equi can be isolated include, in a roughly declining order of frequency29,31–37: (i) Pulmonary (usually the primary site of infection)— sputum, bronchial washings, lung tissue or abscess contents, pleural fluid (empyema) (ii) Blood (predominantly dissemination from lung infection) (iii) Central nervous system (predominantly hematogenous spread)—brain abscesses, cerebrospinal fluid (iv) Subcutaneous and other soft tissue or organ abscesses (predominantly hematogenous spread) (v) Wound drainage, infected joints, vitreous fluid (usually due to traumatic inoculation) (vi) Implanted indwelling devices such as peritoneal dialysis and intravenous catheters (vii) Other sites such as pharynx, middle ear, lymph nodes, bone, and stool. Pulmonary infections are the most common form of human disease caused by R. equi. Cases of lung infection caused by inhalation and cutaneous lesions caused by wound contamination have been documented; the latter are almost the only R. equi infections reported in healthy persons, frequently children.35 In one large review of 72 cases, pneumonia occurred in 76% of patients, and the lung was the sole site of infection in 82% of them.35 Pneumonia was accompanied by extrapulmonary infection in 18% of cases, while

157

Rhodococcus

(a)

(b)

(c)

FIGURE 14.1  (a) Thorax CT-scan and (b) Rx-ray of R. equi bacteremia with lung abscess. (c) Lung histological section with sheets of macrophages with foamy or abundant granular eosinophilic cytoplasm that contain Michaelis-Gutmann bodies, characterized by visible PAS-positive and diastase-resistant basophilic structures with targetoid-like concentric laminations (HE-400×).

infection at extrapulmonary sites occurred without evidence of pulmonary involvement in 24%. Morphology, partial acid fastness, and a distinctive histopathologic profile in bronchial specimens (Figure 14.1c) contribute to accurate diagnosis. Numerous polymorphonuclear leukocytes with intracellular pleomorphic gram-positive bacteria, microabscesses, pseudotumors, and malakoplakia are noted on tissue.29,35 Malakoplakia is a relatively rare granulomatous inflammation not typically associated with histology of lung infection and can be of help in forming a differential diagnosis.35,38,39 There have been few reports of pulmonary infection caused by Rhodococcus equi in immunocompetent hosts. A couple of cases of infection by R. equi in patients without impaired immunity have been reported in the literature, and four of these cases have involved pulmonary disease that showed very different radiological and developmental forms:23,35,40,41 multiple pulmonary nodules and a chronic course; multiple bilateral lobar affectation and acute respiratory failure requiring mechanical ventilation; and atypical community TABLE 14.1 Clinical Manifestations of R. equi Infection as Shown by Various Reports Organ Lung Skin Kidney Bladder Bone Cervix Thyroid Spleen Prostrate glands Colon Liver Heart

Clinical Manifestations Pulmonary nodules, lung abscess, pneumonia, pneumothorax Spontaneous keratitis and endophthalmitis Renal abscess, hepatic abscess Urinary tract infection, bacterimia, isolated and catheter related Septic arthritis, osteomyelitis, infected bone marrow Adenitis along with mesenteric adenitis and peritonitis Thyroid abscess Spleen abscess Prostrate abscess Pedunculated and sessile colonic polyps Hepatic abscess Ventricular shunt infection, sternal-wound infection following coronary bypass grafting

pneumonia. All these patients had cavitation; only one of them required open lung biopsy for diagnosis. Case studies of R. equi infections in human beings indicate diseases related to other organs of the body, too. One study described a kidney transplant recipient who had undergone renal transplantation at the age of 43 years because of endstage renal disease secondary to IgA nephropathy.42 A chest X-ray revealed that the patient had an enlarged left hilus and left-sided pleural effusion. On the patient’s admission to hospital, bronchoscopy findings were macroscopically normal and no mycobacteria were detected in his sputum samples, bronchoalveolar lavage fluid, or pleural fluid. One week after his admission, cultures from these samples and from blood samples returned positive for Rhodococcus equi. The case presented here illustrates the importance of extensive diagnostic procedures in kidney transplant recipients. The clinical presentation of a disease can be atypical in immunocompromised patients, and the spectrum of potential pathologies (e.g., carcinomas) and possible disease-causing microbial agents (e.g., viruses, fungi, and bacteria such as Mycobacterium tuberculosis) in such patients is large. Continued investigations are, therefore, warranted until a definite diagnosis can be made.42 A case study wherein R. equi was the cause of endophthalmitis in an immunocompetent individual showed that the causative agent was a rare R. equi Prescott serotype 4, having been isolated by J. F. Prescott from a few pigs and by the Japanese from horse manure.43 A case of chronic scalp wound infection due to R. equi in an immunocompetent individual following a heavily contaminated traumatic injury was successfully treated by combining antibiotics and surgery.44 This patient was probably exposed to the organism due to the heavy contamination of his wound with soil and his poor personal hygiene. This case helps to elicit the pathogenic potential of R. equi even in the nonimmunosuppressed, as well as to appreciate the variable microbial aetiologies in chronic soft tissue infections and the value of identifying the causative agent. Cases of brain abscesses caused by R. equi have been reported in HIV negative individuals.45,46 In one case, a middle-aged person with a

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history of chronic hepatitis C and alcohol abuse was admitted to the hospital with a 5-day history of hemoptysis and shortness of breath. A magnetic resonance imaging scan of the brain was obtained that showed four ring enhancing lesions in the cerebral cortex. This case emphasizes the role of R. equi in the combined lung-brain abscess syndrome. The study emphasized that all individuals who present with cavitary lung lesions and brain abscesses should be screened for R. equi and treated accordingly. Deaths exceed 50% among AIDS patients with documented R. equi pneumonia and are almost always preceded by multiple relapses, which are common even when successful treatment is achieved.

14.1.3  Pathogenesis and Virulence R. equi infection in animals and in humans show major differences.47 The ability of R. equi to persist in and eventually to destroy macrophages is the basis of its pathogenicity. The pathological change elicited by R. equi is similar to that of some other parasites within the group CorynebacteriumMycobacterium-Nocardia because they have lipid-rich cell wall components.26,48 In nearly all infected foals, the virulence of R. equi is mediated by a surface antigen, VapA, (Virulence-Associated Protein), which is associated with an 85 to 90-kbp plasmid.49,50 Foals infected with VapApositive isolates develop severe bronchopneumonia, whereas plasmid-cured derivatives are harmless. In pigs, nearly all isolates are of intermediate virulence and express a 20-kDa antigen associated with five large plasmids.51 On the other hand, only 20%–25% of isolates recovered from humans express VapA.28,52–54 Expression of the 20-kDa antigen is highly variable and may differ between R. equi strains from different geographical areas. The pathogenesis of infection with intermediately virulent and avirulent strains appears to involve mechanisms different from those associated with VapA-positive isolates.28 The majority of laboratory studies have investigated the virulence and immunity associated with R. equi infection using VapA-positive isolates. Therefore, conclusions about the pathogenesis of R. equi in humans cannot be drawn from these studies. In animals, both humoral and cell-mediated immunity play a role in the control of R. equi infection. Transfer of plasma from immunized horses to infected foals increases opsonization and clearance of R. equi.52 Although normal and CD8_ T cell-deficient mice can clear R. equi infection,55 mice affected with severe combined immunodeficiency (SCID) remain persistently infected but do not develop granulomas.56,57 Nude mice that are given a CD4_ Th1 cell line transferred from immune mice are able to express IFN-g and clear R. equi from their lungs. On the other hand, mice that receive CD4_ Th2 cells but do not receive Th1 cells express IL-4 but not IFN-g, fail to clear the infection, and develop large granulomas with eosinophils in the lung.58,59 Thus, the Th1 response, acting through IFN-g, appears sufficient to effect pulmonary clearance of R. equi while the Th2 response is nonprotective.

Molecular Detection of Human Bacterial Pathogens

This VapA is susceptible to trypsin digestion and accessible to biotin labeling, which suggests that it is present on the outer surface of the cell wall. Radiolabeled VapA was detected as a lipid modified protein anchored to the cell wall.60,61 In gram-positive bacteria, lipoproteins are generally involved in transport of solutes into the cytosol, adherence to surfaces, or have enzymatic activity.62 The signal sequence of lipoproteins contains a “lipobox,” which consists of the amino acid sequence (S/T/G/L/A/V)L(A/S)(G/A)C. The cysteine residue is the first amino acid of the mature protein that is lipid modified when a diacylglycerol moiety is transferred to its sulfhydryl group from a glycerophospholipid molecule. Analysis of the genomic sequence of Mycobacterium tuberculosis revealed the presence of 65 putative lipoproteins containing a lipobox, suggesting that mycolic acid-containing actinomycetes employ a conventional lipid modification system as found in other bacteria.62 However, the VapA protein contains neither cysteine nor a lipobox, suggesting that R. equi may employ an additional mechanism to lipid-modifying proteins. VapA is encoded by the vapA gene that was found to be located on an indigenous R. equi plasmid of approximately 80–85 kb.61,63 Isolates from the submaxillary lymph nodes of infected pigs also contain a large plasmid. However, these plasmids do not encode VapA, but a related protein of 20 kDa (VapB). Interestingly, pig isolates are less virulent in mice than foal isolates harboring the vapA gene, suggesting host specificity of strains harboring different plasmids.51 These studies clearly establish that the virulence plasmid is essential for virulence in foals and pigs, and therefore encodes one or more virulence factors. However, R. equi isolates from cattle and goats lack a virulence plasmid and human isolates contain either VapA or VapB encoding plasmids or lack a virulence plasmid altogether.53,64,65 This suggests that the pathogenesis of R. equi infection in these hosts is different than in foals or pigs. Nucleotide sequencing of the virulence plasmid (Figure 14.2a) of two foal isolates revealed the presence of 69 ORF.66 Based on comparisons with genes previously identified in other organisms three functional regions of the virulence plasmid were recognized. Two of these harbor genes that are similar to those encoding proteins involved in conjugation and in plasmid replication, stability, and segregation. ORF 34, 38, 39, and 40 of the virulence plasmid are similar to the open reading frames found on an indigenous plasmid of Micrococcus sp 28 (Accession number AY034092), and are organized in a similar fashion suggesting a common origin.24 The identification of genes that resemble genes required for conjugation suggest that the virulence plasmid can be mobilized from virulent to avirulent strains. Another segment of ~27.5 kb bears the pathogenicity island (Figure 14.2b) characterized by a significantly lower G + C content than the rest of the plasmid, flanked by genes similar to transposon resolvases, and harbors the vapA gene. Analysis of the pathogenicity island revealed six vapA homologues (vapC, D, E, F, G, H). With exception of vapF, which is not functional, all vap genes encode proteins with a clear signal sequence, indicating that these are extracellular proteins. Byrne et al.

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Rhodococcus

(a)

lssr2 orf60 parB orf63 orf59 resA orf4 orf58 orf5 orf64 vapG orf57 vapH orf56 orf7 orf3 orf8 orf55 orf9 orf54 orf10 orf11 orf53 80610/0 vapA orf52 orf13 parA vapC 10000 70000 trbA vapD orf16 orf 49 orf17 orf 48 orf18 orf 47 vapE p33701 orf 46 60000 20000 vapF 80610 bp orf 45 orf 44 orf21 orf 43 orf 42 invA trbL orf 41 orf23 orf25 orf24 orf 39 30000 50000 orf 38 orf 37 40000 orf 36 orf 35 trsK orf33 orf26 topA orf29 orf31 traA

orf28 orf27 Direct Complement

(b)

ORF1

vapG podG

vapA ORF13 vapC vapD

ORF3

ORFs

ORF4

16–18

ORF5

vapH

vapE

ORFs 7–11

vapF

ORF21

FIGURE 14.2  (a) Map of the R. equi virulence plasmid p33701. The circle shows orientation of the ORFs, with the direction of reading denoted by shading. (b) Schematic view of the 27.5 kb pathogenicity island of the virulence plasmid of R. equi. The arrows indicate the position and direction of transcription of the genes located within the pathogenicity island. White arrows indicate that the genes encode proteins with a signal sequence and may therefore be secreted. Grey arrows indicate the two transcriptional regulators encoded by the pathogenicity island.

showed that VapC, D, and E are secreted, but in contrast to VapA, they are not anchored to the cell wall.67 In addition to the six vap genes, four more genes encode proteins with a clear signal sequence; so actually there are 10 total extracellular proteins encoded by the pathogenicity island. As extracellular proteins can interact with the host environment, these 10 proteins could be important virulence factors. ORF8 encodes a response regulator, which usually interacts with a sensor kinase protein to regulate gene expression. The later autophosphorylates conserved histidine residue in response to an environmental signal; the phosphate is transferred to a conserved aspartate residue in the response regulator resulting in transcriptional activation. Although the virulence plasmid encodes a response regulator, it does not encode a sensor kinase, indicating that the ORF8 gene product interacts

with a chromosomally encoded sensor kinase protein. The PhoP/PhoQ system is an example of a two-component system in Salmonella typhimurium that is induced within the phagosome of the macrophage.68 This regulatory system has also been identified in Yersinia pestis and was shown to be important for survival under conditions of macrophage-induced stress.69 ORF4 encodes a protein homologous to the LysR type transcriptional regulators (LTTR). LTTR are, after the twocomponent response regulators, the second largest class of bacterial regulatory proteins.70 A number of LTTR have been identified that are associated with virulence gene regulation. For example, SpvR encoded by the 90 kb virulence plasmid of Salmonella typhimurium, regulates the spv operon in response to conditions such as late growth phase.71 Interestingly, the spv operon has been shown to be induced in bacteria within

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macrophages.72 Three other genes, ORF 3, 5, and 21, encode proteins with clear homologues in other bacteria. The protein encoded by ORF 21, which has a M. tuberculosis homologue (Rv1885c), is a secreted chorismate mutase, an enzyme of the shikimate pathway for biosynthesis of aromatic amino acids. The extracellular location of this protein is enigmatic, as this pathway is usually located inside the cytoplasm. The protein encoded by ORF 5 is predicted to be an integral membrane protein, displaying a strong similarity to sugar permeases (Pfam 00083). ORF 3 encodes an O-methyl-transferase protein that lacks a signal sequence and is therefore likely to be located in the cytoplasm. Although it has not yet been shown whether these genes are required for virulence, their presence in the pathogenicity island might suggest that virulent R. equi either produces a metabolite that is secreted by the cell or that R. equi takes up a modified host metabolite. The former could serve to alter the macrophage response; the latter could be involved in signaling. Studies show that the regulation of expression of genes within the pathogenicity island is complex, and depends on at least five environmental signals: temperature, pH, oxidative stress, magnesium, and iron. It seems very likely that the two transcriptional regulators encoded by ORF4 and ORF8 play an important role in transducing these signals to the transcription apparatus, although this has not yet been established. To date, two transcriptional repressors, Fur and IdeR, controlling iron-dependent gene expression have been identified.73 An ideR homologue of R. equi was recently cloned and overexpressed in E. coli. It was shown that the R. equi IdeR protein is functional; it binds to a consensus IdeR recognition sequence and regulates gene expression in an Fe2+ dependent manner.74 Interestingly the promoter region of the vapA gene contains an IdeR consensus binding site,75 suggesting that the iron-dependent expression of this gene may be controlled by IdeR.

14.1.4  Cell Biology It is understood that nonspecific degranulation of lysosomes in R. equi-infected macrophages was suggested to be the major survival and pathogenic mechanism of infection in horses.76 Such degranulation in vivo would contribute to the tissue destruction and neutrophil influx that is characteristic of R. equi lung lesions.76 The basis of pathogenicity of R. equi is its ability to multiply in and eventually to destroy alveolar macrophages. Since opsonization of R. equi with specific antibody is followed by increased phagosome-lysosome fusion and by significantly enhanced R. equi killing by equine macrophages,76,77 macrophage entry through non-Fc receptors may be important in determining the fate of the bacteria. Once bacteria are internalized, the phagosome undergoes a series of fusion and fission events with vesicles from the endocytic pathway, a complex maturation process that leads to the formation of phagolysosomes. During maturation, the phagosome membrane increasingly changes to resemble that of late endosomes and lysosomes. For bacteria that do not interfere with the process, phagocytosis to phagolysosome formation

Molecular Detection of Human Bacterial Pathogens

takes about 5 min.78 Phagocytosed bacteria are degraded in the late phagolysosome by the same mechanisms employed by lysosomes, involving a wide range of acid-resistant hydrolases within the acid environment of the phagolysosome. Intramacrophage R. equi appear to be located exclusively within membrane-enclosed vacuoles and persistence correlates with the absence of phagosome-lysosome fusion.76 It has been shown that plasmid-encoded products contribute to the ability of R. equi to survive and replicate in macrophages. Preliminary observations suggested that maturation of the phagosome is more efficiently diverted in strains possessing the virulence plasmid compared to plasmid-negative isogenic strains, and that cytotoxicity of R. equi for J774E murine macrophages is strongly upregulated by the possession of the virulence plasmid.24

14.1.5  Diagnosis 14.1.5.1  Phenotypic Identification Isolation and Colonial Morphology. R. equi grows readily when incubated aerobically at 37°C on nonselective media routinely used in clinical microbiology laboratories. At 24 h of incubation, colonies are 1–2 mm in diameter and are not distinctive. By 48 h of incubation on nonselective media such as Trypticase soy blood agar, they developed their characteristic appearance26 exhibited by irregularly round, smooth, semitransparent, glistening, coalescing, mucoid, teardrop colonies with entire edges. The colonies vary in size from 2 to 4 mm, although coalesced colonies may appear larger. The organism does not grow on most types of MacConkey agar. Rare isolates of R. equi may grow poorly at 37°C.79 Colony variation is present in fresh cultures. The classical viscous-mucoid coalescing colony is usually the predominant type present, but less mucoid forms may also be seen. Cultures may have a slightly earthy smell. Pigment production is rarely marked in cultures that are less than 4 days old80 and it may surprise those expecting all rhodococci (red-pigmented cocci) to show obviously pink or red colonial pigmentation immediately on isolation. After 4–7 days on nonselective medium such as Trypticase soy blood agar, colonies may develop a delicate shade of salmon pink, although they may be nonpigmented or appear slightly yellow.26,80 Perhaps the best description of the typical colony pigment on blood agar is as light fawn colored. Cultures kept on slopes for prolonged periods without subculture commonly become rough, dry, and orange-red but revert to classical colonies on subculture.81 The macroscopic colony morphology of the type culture ATCC 693981 is pigmented than typical R. equi. Isolation of R. equi from the infection site is so far the best method of diagnosing infection. Sputum is an often but not always useful specimen in humans with pneumonic disease.82 Diagnosis is more reliably made by bronchial brushings, percutaneous thoracic aspiration, or, more drastically, open lung biopsy during lobectomy.83 On occasion, bronchial brushings in human patients have failed to yield R. equi.83 This failure supports the findings of Martens et al. that culture of bronchial secretions obtained by transtracheal aspiration from

161

Rhodococcus

experimentally infected foals does not consistently yield R. equi.84 They attributed this inconsistency to either periodic shedding into airways or the inability of organisms inside phagocytes to be isolated by standard culture techniques. Although most culture-positive foals with R. equi pneumonia yield pure cultures of organisms from lung aspirates, up to one-third may also yield a variety of opportunist pathogens.26 Thus, R. equi may be present in mixed culture and may be ignored in part because it does not show its distinctive colony size and mucoid appearance until after 48 h of incubation. Blood cultures from febrile human patients with pneumonic disease have yielded R. equi (sometimes repeatedly) in about one-third of cases reported.85 Microscopic Studies and Staining Properties. R. equi is a gram-positive pleomorphic coccobacillus that varies from distinctly coccoid to bacillary depending on growth conditions. It is usually coccoid, both on solid media and in purulent material from patients, but in liquid media, particularly in young cultures, it forms long rods or short filaments that may show rudimentary branching. The occasional reports that R. equi is acid fast in the Ziel–Neelsen stain appear to depend on staining technique, the older age of the cultures, and the growth medium.26,86 Most report a failure to demonstrate acid fastness of R. equi. Metachromatic granules have occasionally been described,26 but like acid-fast staining, this is an inconsistent feature of the organism. Biochemical Characterization. Characteristics that can be used in routine clinical microbiology laboratories to identify R. equi are shown in Table 14.2. The organism is generally biochemically unreactive. R. equi is an obligate aerobe TABLE 14.2 Biochemical Characteristics for Identifying R. equi in Comparison to Corynebacterium and Other Closely Related Pathogens Characteristic

Reaction

Beta-hemolysis Esculin hydrolysis Nitrate reduction Urease Casein digestion Glucose Maltose Sucrose Catalase Cytochrome coxidase Carbohydrate fermentation Alcohol fermentation Gelatin hydrolysis Indole H2S Hippurate hydrolysis Nitrate reduction equi factors (CAMP test) Lipase Phosphatase

− − + + − − − − + − − − − − Variable − + + + +

with simple growth requirements. It can utilize carbon from a wide variety of sole carbon sources, including acetic, pyruvic, butyric, and propionic acids, and nitrogen from ammonium sulfate or potassium nitrate.26 The organism is nonmotile and nonflagellated and may produce small numbers of pili.86 Some strains may produce a thin pellicle that is readily disrupted. It fails to oxidize or ferment most carbohydrates or alcohols and is nonproteolytic. No acid metabolic products from glucose were detected by gas-liquid chromatography.87 It is catalase positive, almost invariably urease positive, and oxidase negative. R. equi produces equi factors that interact with the phospholipase D of Corynebacterium pseudotuberculosis, the beta-toxin of S. aureus, and an uncharacterized partial hemolysin of L. monocytogenes to give an area of complete hemolysis with sheep or cattle erythrocytes. While this characteristic is not unique for R. equi, it is distinctive for this organism and should always be used in identification since no equi factor-negative isolates of R. equi have been described. Antimicrobial drug susceptibility is consistent and might be a useful adjunct in identification. R. equi produces lipase and phosphatase, but not DNase, elastase, lecithinase, or protease. The API ZYM (Analytab Products Inc., Plainview, NY) gave distinctive enzyme profiles of value in identifying R. equi. All isolates tested were positive for leucine arylamidase, acid phosphatase, and phosphamidase; >90% were positive for valine amylamidase, esterase lipase, and alpha-glucosidase.26 Capsular Serotypes. R. equi possesses a distinct, antigenically variable, lamellar polysaccharide capsule, which has provided the basis of a capsular serotyping system.88 At least 27 different capsular serotypes have been identified, of which Prescott capsular serotype 1 (or its equivalent in other typing schemes) is the most common worldwide.26,88 No relationship between serotype and virulence is apparent. A number of workers have investigated alternative, immunological approaches to diagnosis in foals but none of those can be recommended as replacing cultural approaches. In immunosuppressed human patients, including those with AIDS, immunological tests cannot be depended on for diagnosis.26 Pathologic findings in patients with R. equi infection are relatively consistent, despite the broad range of tissues involved. Specimens are typically necrotic and contain a dense, histiocytic infiltrate with an eosinophilic, granular cytoplasm, and intrahistiocytic coccobacilli. Multiple microabscesses that contain abundant neutrophils may be present. In some cases, concentrically layered basophilic inclusions, called “Michaelis–Gutman bodies,” are prominent. The inclusions are thought to result from impaired macrophage lysosomal function that leads to failure of lysophagosomal fusion and ineffective killing of ingested organisms.89 This characteristic pathologic finding, together with Michaelis-Gutman bodies, is termed “malakoplakia,” and has been described in other bacterial infections.29,47 In rare cases, a dense proliferation of spindle cells can mimic either Kaposi’s sarcoma or lymphosarcoma.29,47 A synergistic hemolysis-inhibition assay for antibody to equi factors distinguished foals with naturally occurring

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R. equi pneumonia from healthy foals, although it failed to identify foals in the early stages of experimentally induced disease.90 An agar gel immunoprecipitation test for antibody to equi factors was also shown to identify foals with R. equi pneumonia, although a small proportion of apparently healthy foals as also positive in the test.26 Using more sensitive synergistic-hemolysis inhibition and immunoprecipitation assays than described previously, Skalka demonstrated the widespread nature of subclinical infection or exposure to R. equi in foals.26,91 In general, ELISA has been used to distinguish infected from healthy foals.92 In one study, a Tween 20 extract from the type strain, ATCC 6939, gave the broadest cross reactivity of the serotypes tested.26,92 The mean values of anti-R. equi IgG antibody in foals known or suspected to be ill with R. equi pneumonia were strikingly higher than in normal foals in the limited number of animals tested, but there were no differences in mean values of anti-R. equi IgM or IgA antibodies.92 This ELISA was more sensitive than agar gel diffusion or indirect hemagglutination tests.92 In two unrelated studies, using different antigens, no correlation was found between ELISA antibody levels and clinical disease.26 The antigens used in ELISA are important in determining their value for serological diagnosis.92 Supernatants of autoclaved cells or of homogenized cells were relatively poor antigens compared with Tween 20-extracted or sonicated materials.92 Two unique features of R. equi help distinguish it from other organisms. First, synergistic hemolysis occurs when R. equi cultured on sheep blood agar are cross-streaked with other bacteria, including Staphylococcus aureus, Listeria monocytogenes, and Corynebacterium pseudotuberculosis.26 Second, antagonism between imipenem and other β-lactam antibiotics has been documented in vitro. This antagonism is widespread among R. equi isolates and is novel among grampositive bacteria.47,93 Serologic assays have not been clinically validated and are not commercially available. 14.1.5.2  Molecular Identification Polymerase chain reaction (PCR) detection of R. equi can offer rapid detection of this pathogenic bacterium, however, only a few rapid molecular methods have been developed for R. equi identification. Clinical isolates from foals can be identified by detection of the VapA antigen with monoclonal antibodies, or alternatively, by PCR detection of its gene, vapA, which is present on an 85 kb virulence plasmid.66,94,95 One such study has shown that PCR of tracheal wash using primers that recognized the vapA virulence plasmid of R. equi has a diagnostic sensitivity of 100% and specificity of 90.6%. In contrast, sensitivity and specificity are only 57.1% and 93.8%, respectively, for standard microbiologic culture of tracheal wash and only 62.5% and 75.9%, respectively, for serology.96 However, the virulence plasmid is not present in all strains of human and environmental origin (in general, in nonequine isolates),53 thus limiting the general usefulness of the vapA-based identification of R. equi by PCR. In another study, species-specific primers for the PCR for the detection of Rhodococcus equi were developed. These primers were based on unique DNA fragments produced from R.

Molecular Detection of Human Bacterial Pathogens

equi reference strains and field isolates. Following random amplification of polymorphic DNA from R. equi and R. rhodochrous with a set of 40 arbitrary 10 bp primers, a pair of species-specific primers was designed to detect a unique 700 bp fragment of R. equi chromosomal DNA. This PCR product was limited to R. equi and was not detectable in other Rhodococcus species or in a panel of additional gram-positive and gram-negative bacteria.97 Other PCR-based molecular methods have been used to amplify the 16S rRNA,98–100 but their validities have been assessed with only a small number of strains. Although 16S rRNA sequencing is accepted as a general means for species differentiation, some heterogeneity can exist between different isolates of the same species.100 A recent study of the 16S rRNA sequences of several representative strains showed that R. equi is a very heterogeneous taxon, with variations in 16S rRNAs of up to 4%.101 On the other hand, closely related species may have identical or almost identical 16S rRNA sequences.102 Therefore, it is useful to have other species-specific targets, such as choE, to undertake an assay for the reliable identification of a bacterial species. Ladron et al. reported a specific PCR assay for the rapid and reliable identification of R. equi based on the amplification of a fragment of the choE gene encoding cholesterol oxidase. The choE-based PCR was assessed by using a panel of strains comprising 132 isolates from different sources and of different geographical origins, all initially identified biochemically as R. equi, and 30 isolates of representative nonR. equi actinomycete species, including cholesterol oxidase producers. The expected 959 bp amplicon was observed only with R. equi isolates, as confirmed by sequencing of a variable region of the 16S rRNA gene from a random sample of 20 PCR-positive isolates. All R. equi isolates gave a positive choE-based PCR result, which correlated with a high degree of conservation of the choE gene.103 Finally, a PCR-RFLP method targeting a 65-kDa heat shock protein gene was shown to discriminate R. equi strains.104 This test was primarily devised for the identification of mycobacteria. However, this assay is too laborious and lengthy because, due to the conserved nature of the 65-kDa heat shock protein gene, the amplification product is of the same size for all actinomycete species and R. equi can be discriminated only by restriction analysis of the amplicon. Advanced techniques, such as multiplex PCR was used for molecular characterization of this bacterium for epidemiological studies.105,106 Therefore, these studies suggest that the molecular detection of this pathogen is an area of current research interest.

14.1.6  Treatment and Control Treatment. A proposed approach for the treating patients with Rhodococcus equi infections based on in vitro susceptibility data and published case reports47 has been outlined in Figure 14.3. These infections can be successfully treated using lipophilic antimicrobial drugs that can penetrate the macrophages or neutrophils in which the organisms survive.26,107 A number of in vitro studies are being carried out to study the

163

Rhodococcus

Patient Localized infections

Disseminated clinical signs

Culturing and susceptbility testing Oral therapy

Trachypea Fever Cough Nasal discharge

Intravenous therapy

Confirm anitimicrobial regimen concurs with results of susceptibility testing R. equi isolated or at least 2 attached conditions If yes Oral therapy Repeat cultures Consider chronic suppression if immunocompetent or central nervous system infection

• Multifocal pulmonary opacities on radiographs • Pulmonary abscesses on ultrasound • Gram positive intracellular coccobacilli on TBA specimens (culture plate)

Terminate therapy if symptoms and radiologic abnormalities are resolved

FIGURE 14.3  Proposed approach for the treatment of R. equi infection. Intravenous antibiotics are typically required for initial treatment, except in the case of localized infection in an immunocompetent host. Antimicrobial agents are rated as “preferred” or “alternative” on the basis of their in vitro activity, the results of animal studies, clinical experience, and published case reports.

antimicrobial susceptibility of clinical isolates since they cause life-threatening infections in severely immunocompromised persons.43,108,109 While variations exist, most strains were susceptible to inhibition by glycopeptide antibiotics (including vancomycin and teicoplanin) and rifampin. Macrolide antibiotics, such as erythromycin and clarithromycin, were also inhibitory to many strains. Resistance to β-lactam antibiotics (with the exception of carbapenems, specifically imipenem) was generally reported, and is not related to the production of a β-lactamase. The standard treatment in pneumonic foals is oral administration of a combination of erythromycin estolate and rifampin.110 These drugs have also been used alone or in combination, usually with good clinical response, in many human infections.26,111,112 Unfortunately, however, there is no standard treatment protocol for pulmonary and/or systemic R. equi infections as relapse in spite of treatment is common in a majority of cases,108 followed by high mortality rate, especially among AIDS patients (Table 14.1). A proposed regimen involves parenteral glycopeptide plus imipenem for at least 3 weeks, followed by an oral combination of rifampin, plus either macrolides or tetracycline.43 Examples of efficacious protocols for parenteral treatment are available (Table 14.1). Susceptibility testing should be performed for all R. equi isolates, to avoid the use of antibiotics to which the isolate has in vitro resistance. R. equi is usually susceptible in vitro to a number of substances, including erythromycin, rifampin, flouroquinolones, aminoglycosides, glycopeptides, and imipenem. Susceptibility to cotrimoxazole, tetracycline, chloramphenicol, clindamycin, and cephalosporins is variable. Isolates are typically resistant to penicillins, but use of penicillins is not

recommended because of rapid acquisition of resistance.12,35,113 For immunocompromised patients and patients with serious infections, intravenous therapy with two-drug or three-drug regimens that include vancomycin, imipenem, aminoglycosides, ciprofloxacin, rifampin, and/or erythromycin are appropriate. Regimens that include 12 drugs have been successful for some patients, but no data are available to indicate that they are superior. Most patients will require intravenous antibiotics for a minimum of 2 weeks, at which point clinical improvement should be evident. Oral antibiotics can then be substituted and continued until all culture results are negative and the patient’s symptoms and radiologic abnormalities have resolved.26,114 The duration of therapy depends on the site(s) and extent of infection, underlying immunocompetence of the host, and the clinical response to therapy. A minimum of 6 months of antibiotic therapy is typically required for immunocompromised patients. Besides the use of antimicrobial drugs, the approach used in treatment of human infection involves drainage of suppurative lesions, surgical resection of granulomatous tissue, and control of predisposing factors such as decreasing the dose of concurrent immunosuppressive drugs or control of underlying malignancies. Careful and repeated culture and susceptibility testing during treatment is required to discover acquired resistance, in a manner similar to the treatment of mycobacterial infection. Prevention and Control. Primary prophylaxis against Rhodococcus equi is not routinely recommended, because sufficient data is not available to support its efficiency and also the infection is rare. Macrolide prophylaxis against

164

Mycobacterium avium complex infection may offer some protection against R. equi infection for patients with AIDS. Immunocompromised patients with significant exposure to domesticated animals should be cautioned regarding the possible risk of R. equi infection. Some investigators have advocated isolation of hospitalized patients with R. equi pneumonia to prevent nosocomial spread.115,116 There is no effective vaccine currently available for the prevention of R. equi pneumonia. DNA vaccines are known to offer specific advantages over conventional vaccines. Some recombinant DNA vaccine candidates, namely pcDNA3-Re1, pcDNA3-Re3, and pcDNA3-Re5, were designed by combining a heat shock protein GroEL2 to a virulence-associated protein A (VapA) from R. equi to protect C3H/He mice against the R. equi infection. VapA was shown to be strongly recognized by sera from pneumonic foals. All vaccines elicited at least a doubling of the IgG2a/IgG1 ratio in comparison to the controls, indicating a bias to the Th1 response, which is postulated to be crucial for bacterial clearance and protective immunity against intracellular pathogens, including R. equi.117 Improved methods to effectively control this pathogen and treat infections caused by it are areas of current research and holds potential challenges.

Molecular Detection of Human Bacterial Pathogens

sensitivity and 93.8% specificity found respectively in the standard microbiologic culture procedures of tracheal wash. For serology, the sensitivity and specificity were found to be only 62.5% and 75.9%, respectively. The PCR protocol described here was originally reported by Ladron et al.103:





14.2  METHODS 14.2.1  Sample Preparation DNA can be extracted from tracheal aspirate samples of infected foals as follows:118 A volume of 300 μL of sodium iodide solution containing 6 M NaI, Tris hydrochloride (pH 8), 13 mM EDTA, 0.5% sodium N-lauroylsarcosine, and 10 μg of glycogen is added to tubes, each containing 100 μL of tracheal aspirate sample. The tubes are incubated at 60°C for 15 min. A volume of 400 μL of 100% isopropanol is added to each tube, mixed thoroughly, and left at room temperature for 15 min. The tubes are centrifuged at a speed of 10000 × g for 15 min and the supernatant is completely discarded. A volume of 1 mL of Washing Solution B is used to suspend the pellet in each tube, followed by vigorous vortexing. Each tube is briefly centrifuged at 10000 × g for 5 min. DNA is resuspended in TE buffer after the completely discarding supernatant from each tube. This protocol is actually the DNA extraction protocol followed in the manufacturer’s instructions of the DNA Extractor kit produced by Wako Pure Chemical Industries, Japan.

(ii) Restriction Fragment Length Polymorphism (RFLP) Analysis  The PCR products may be purified by use of the Gel Extraction Purification Kit (Qiagen) and subsequently digested with the following restriction enzymes for generating restriction patterns: AvaI, BamHI, BglI, HinPI, PvuI, XhoI, and XmnI (New England Biolabs). The digestion products are resolved by electrophoresis on 1.2% agarose gels in Tris borate buffer and visualized by ethidium bromide staining. (iii) Multiplex PCR Assay for the vap Gene Family  The primers for PCR amplification for vap genes were designed with Vector NTI 8.0 software (InforMax Inc., 2002) based on sequences published through GenBank.119 PCR amplification is performed as described previously118 with some modifications.

14.2.2  Detection Procedures (i) PCR  PCR is a very efficient tool to detect rapidly the presence of R. equi. Sellon et al.96 demonstrated that detection of R. equi by performing PCR on tracheal wash samples of infected foals using primers that recognized the vapA virulence plasmid of R. equi is 100% more sensitive and 90.6% more specific, as compared to the 57.1%

1. The oligonucleotide primers are synthesized flanking the segment to be amplified. 2. PCR mixture is prepared by mixing 10 μL of template, 40 pmol of each oligonucleotide primer, 4 U of Taq DNA polymerase (Bioline, London, United Kingdom), 0.2 mM each deoxynucleoside triphosphate, 1.5 mM MgCl2, and 10 μL of 10× PCR amplification buffer [160 mM (NH4)2SO4, 670 mM Tris-HCl (pH 8.8), 0.1% Tween 20]. Finally, doubledistilled water is added to the PCR mixture to make a final volume of 100 μL. 3. Run the following cycling program in a PCR machine: one cycle of 95°C for 5 min; 30 cycles of 95°C for 1 min, 55°C for 1 min, and 72°C for 1 min; and one cycle of 72°C for 10 min. 4. The amplified DNA is visualized generally on a 0.7% agarose gel by UV fluorescence (approximately 5 μL sample), following staining with ethidium bromide.118



1. For each reaction 4 µL (50 ng) of DNA preparation is used in a 25 µL reaction mixture containing 10 mM Tris, 50 mM KCl, 2.5 mM MgCl2, 200 mM (each) deoxynucleoside triphosphates, 1 µM of each primer (30 pmol), and 1 U of Taq DNA polymerase. 2. The PCR mixture is subjected to 30 cycles of 94°C of 90 s, 55°C for 1 min, and 72°C for 2 min, with a final extension at 72°C for 10 min. 3. The amplification products (15 µL) is separated by 1.5% agarose gel electrophoresis for 60 min at 80 V. DNA fragments are visualized by UV fluorescence after staining with ethidium bromide.

165

Rhodococcus

TABLE 14.3 Shuttle Vectors for Use in R. equi and Other Species Name pRE-1 pRE-7 pREV5

pMS21 pJW1484 pLG1484

Plasmids Used in Constructing Vector pACYC177 (E. coli) pOTS (ori from R. equi) pRE-1 pBS (vapA subcloned in MCS) pAL5000, pMF1 (ori from Several E. coli vectors and M. fortuitum) pHP45X-apra pMF1 (ori from M. fortuitum) pSUM36 and pEP2 (E. coli) pMS1 (Corynebacterium strain) pBR322 (E. coli) pMS2 pE194 (S. aureus and B. subtilis) pMS2 pTV543 (B. subtilis) pIAM1484 (R. erythropolis) pBSk (-) E. coli pJW1484 was modified for higher replication stability

pDA37

Nocardioform plasmid

pREV2 pMS2 pMS20

E. coli-positive selection vector constructed by M. Zabeau et al.

(iv) Sequencing  Control strains (without vap genes) may be sequenced in an automatic auto sequencer (MegaBace 500). Each vap gene should be separately submitted for PCR and the products obtained may be purified with polyethyleneglycol (PEG 8000). The sequences can be analyzed by submitting them to a consensus analysis based on chromatogram reliability obtained using the Staden Package Gap 4 program.120 Thereafter, samples may be compared to vap gene sequences stored at GenBank using the Chromas Proprogram, version 1.34.

Use Regulation of virulence gene expression in R. equi These vectors were compatible in R. equi. Used for analysis of gene functions in trans Study gene function in a wide range of bacterium: R. equi, Corynebacterium, E. coli, B. subtilis, S. aureus Relatively small in size, can replicate in a variety of species of Rhodococcus and has high replication stability in a host Cloning DNA without prior alkaline phosphatase treatment

Reference 121 122

Katsumata, R. et al., U.S. Pat. No. 4,500,640 United States Patent Application 20060263868 123

organism have been successful in preventing pneumonia in mice, although further studies are required to develop effective vaccines in foals and humans. Molecular detection of the pathogen in clinical samples and knowledge on molecular basis of pathogenesis and epidemiology are also primary areas of future research work. In recent times, due to the upsurge of various molecular techniques, study of such diseases has become potentially easier and quicker. Therefore, foolproof vaccines and other means of cure can be anticipated to be available soon.

ACKNOWLEDGMENTS 14.3  C  ONCLUSION AND FUTURE PERSPECTIVES Much has been learned in the last decades about R. equi infections in foals that can be applied to the diagnosis and treatment of infection in humans. R. equi is a rare but recognized pathogen in humans and has emerged as an important cause of morbidity and mortality among immunocompromised patients. Increasing awareness of R. equi infection improves the likelihood of its accurate and timely diagnosis. Although considerable progress has been made in understanding important aspects of virulence of R. equi, there is an almost complete lack of genetic tools for this bacterium that will be required to make further progress. To address this problem R. equi–E. coli shuttle vectors have been developed to facilitate transformation and random as also site-specific mutagenesis procedures (Table 14.3). The shuttle vectors that replicate in R. equi are based on different replicons so that they can be maintained in the cell at the same time. Random and transposon-based site directed mutagenesis techniques have also been applied to study virulence of this pathogen. There is no effective vaccine currently available for the prevention of R. equi pneumonia. Chimeric DNA vaccines are being formulated and tested for their efficiency in mice. Vaccines targeting the vapA and heat shock proteins117 in this

We thank Dr. Debdas Mukherjee and Dr. Biswajit Paul for useful suggestions. Debarati Paul is grateful to Dr. Mark Lawrence and Dr. Shane Burgess for their support.

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167 68. Alpuche Aranda, C.M. et al., Salmonella typhimurium activates virulence gene transcription within acidified macrophage phagosomes, Proc. Natl. Acad. Sci. USA, 89, 10079–10083, 1992. 69. Oyston, P.C. et al., The response regulator PhoP is important for survival under conditions of macrophage-induced stress and virulence in Yersinia pestis, Infect. Immun., 68, 3419–3425, 2000. 70. Schell, M.A., Molecular biology of the LysR family of transcriptional regulators, Annu. Rev. Microbiol., 47, 597–626, 1993. 71. Caldwell, A.L., and Gulig, P.A., The Salmonella typhimurium virulence plasmid encodes a positive regulator of a plasmidencoded virulence gene, J. Bacteriol., 173, 7176–7185, 1991. 72. Rhen, M., Riikonen, P., and Taira, S., Transcriptional regulation of Salmonella enterica virulence plasmid genes in cultured macrophages, Mol. Microbiol., 10, 45–56, 1993. 73. Hantke, K., Iron and metal regulation in bacteria, Curr. Opin. Microbiol., 4, 172–177, 2001. 74. Boland, C.A., and Meijer, W.G., The iron dependent regulatory protein IdeR (DtxR) of Rhodococcus equi, FEMS Microbiol. Lett., 191, 1–5, 2000. 75. Ren, J., and Prescott, J.F., Analysis of virulence plasmid gene expression of intra-macrophage and in vitro grown Rhodococcus equi ATCC 33701, Vet. Microbiol., 94, 167– 182, 2003. 76. Hietala, S.K., and Ardans, A.A., Interaction of Rhodococcus equi with phagocytic cells from R. equi-exposed and nonexposed foals, Vet. Microbiol., 14, 307–320, 1987. 77. Zink, M.C. et al., Electron microscopic investigation of intracellular events after ingestion of Rhodococcus equi by foal alveolar macrophages, Vet. Microbiol., 14, 295–305, 1987. 78. Mayorga, L.S., Bertini, F., and Stahl, P.D., Fusion of newly formed phagosomes with endosomes in intact cells and in a cell-free system, J. Biol. Chem., 266, 6511–6517, 1991. 79. Muller, F. et al., Characterization of Rhodococcus equi-like bacterium isolated from a wound infection in a noncompromised host, J. Clin. Microbiol., 26, 618–620, 1988. 80. Elissalde, G.S., Renshaw, H.W., and Walberg, J.A., Corynebacterium equi: An interhost review with emphasis on the foal, Comp. Immunol. Microbiol. Infect. Dis., 3, 433–445, 1980. 81. Williams, G.D., Flanigan, W.J., and Campbell, G.S., Surgical management of localized thoracic infections in immunosuppressed patients, Ann. Thorac. Surg., 12, 471–482, 1971. 82. Golub, B., Falk, G., and Spink, W.W., Lung abscess due to Corynebacterium equi. Report of first human infection, Ann. Intern. Med., 66, 1174–1177, 1967. 83. Savdie, E., Pigott, P., and Jennis, F., Lung abscess due to Corynebacterium equi in a renal transplant recipient, Med. J. Aust., 1, 817–819, 1977. 84. Martens, R.J., Fiske, R.A., and Renshaw, H.W., Experimental subacute foal pneumonia induced by aerosol administration of Corynebacterium equi, Equine Vet. J., 14, 111–116, 1982. 85. Berg, R. et al., Corynebacterium equi infection complicating neoplastic disease, Am. J. Clin. Pathol., 68, 73–77, 1977. 86. Yanagawa, R., and Honda, E., Presence of pili in species of human and animal parasites and pathogens of the genuscorynebacterium, Infect. Immun., 13, 1293–1295, 1976. 87. Reddy, C.A., and Kao, M., Value of acid metabolic products in identification of certain corynebacteria, J. Clin. Microbiol., 7, 428–433, 1978.

168 88. Nakazawa, M.K. et al., Serogrouping of Rhodococcus equi, Microbiol. Immunol., 27, 837–846, 1983. 89. Drancourt, M. et al., Rhodococcus equi infection in patients with AIDS, J. Infect., 24, 123–131, 1992. 90. Prescott, J.F., Coshan-Gauthier, R., and Barksdale, L., Antibody to equi factor(s) in the diagnosis of Corynebacterium equi pneumonia of foals, Can. J. Comp. Med., 48, 370–373, 1984. 91. Skalka, B., and Svastova, A., Two techniques for detection of antibodies against Corynebacterium (Rhodococcus) equi in horse sera, Vet. Microbiol., 10, 293–300, 1985. 92. Takai, S., Kawazu, S., and Tsubaki, S., Enzyme-linked immunosorbent assay for diagnosis of Corynebacterium (Rhodococcus) equi infection in foals, Am. J. Vet. Res., 46, 2166–2170, 1985. 93. Nordmann, P., Nicolas, M.H., and Gutmann, L., Penicillinbinding proteins of Rhodococcus equi: Potential role in resistance to imipenem, Antimicrob. Agents Chemother., 37, 1406–1409, 1993. 94. Takai, S. et al., Evaluation of a monoclonal antibody-based colony blot test for rapid identification of virulent Rhodococcus equi, J. Vet. Med. Sci., 56, 681–684, 1994. 95. Takai, S. et al., Identification of virulent Rhodococcus equi by amplification of gene coding for 15- to 17-kilodalton antigens, J. Clin. Microbiol., 33, 1624–1627, 1995. 96. Sellon, D.C. et al., Comparison of nucleic acid amplification, serology, and microbiologic culture for diagnosis of Rhodococcus equi pneumonia in foals, J. Clin. Microbiol., 39, 1289–1293, 2001. 97. Arriaga, J.M. et al., Detection of Rhodococcus equi by polymerase chain reaction using species-specific nonproprietary primers, J. Vet. Diagn. Invest., 14, 347–353, 2002. 98. Bell, K.S. et al., Identification of Rhodococcus equi using the polymerase chain reaction, Lett. Appl. Microbiol., 23, 72–74, 1996. 99. Sellon, D.C. et al., Nucleic acid amplification for rapid detection of Rhodococcus equi in equine blood and tracheal wash fluids, Am. J. Vet. Res., 58, 1232–1237, 1997. 100. Kirschner, P., and Bottger, E.C., Microheterogeneity within rRNA of Mycobacterium gordonae, J. Clin. Microbiol., 30, 1049–1050, 1992. 101. McMinn, E.J. et al., Genomic and phenomic differentiation of Rhodococcus equi and related strains, Antonie Van Leeuwenhoek, 78, 331–340, 2000. 102. Patel, R. et al., Determination of 16S rRNA sequences of enterococci and application to species identification of nonmotile Enterococcus gallinarum isolates, J. Clin. Microbiol., 36, 3399–3407, 1998. 103. Ladron, N. et al., Rapid identification of Rhodococcus equi by a PCR assay targeting the choE gene, J. Clin. Microbiol., 41, 3241–3245, 2003. 104. Taylor, T.B. et al., Routine use of PCR-restriction fragment length polymorphism analysis for identification of mycobacteria growing in liquid media, J. Clin. Microbiol., 35, 79–85, 1997. 105. Ocampo-Sosa, A.A. et al., Molecular epidemiology of Rhodococcus equi based on traA, vapA, and vapB virulence plasmid markers, J. Infect. Dis., 196, 763–769, 2007.

Molecular Detection of Human Bacterial Pathogens 106. Krewer, C.D.C. et al., Molecular characterization of Rhodococcus equi isolates of horse breeding farms from an endemic region in South of Brazil by multiplex PCR. Vol. 39 188–193 (scielo, 2008). 107. Hillidge, C.J., Review of Corynebacterium (Rhodococcus) equi lung abscesses in foals: Pathogenesis, diagnosis and treatment, Vet Rec., 119, 261–264, 1986. 108. Bernheimer, A.W., Linder, R., and Avigad, L.S., Stepwise degradation of membrane sphingomyelin by corynebacterial phospholipases, Infect. Immun., 29, 123–131, 1980. 109. Eales, L.J., and Parkin, J.M., Current concepts in the immunopathogenesis of AIDS and HIV infection, Br. Med. Bull., 44, 38–55, 1988. 110. Hillidge, C.J., Use of erythromycin-rifampin combination in treatment of Rhodococcus equi pneumonia, Vet. Microbiol., 14, 337–342, 1987. 111. Hillerdal, G. et al., Infection with Rhodococcus equi in a patient with sarcoidosis treated with corticosteroids, Scand. J. Infect. Dis., 20, 673–677, 1988. 112. Samies, J.H. et al., Lung abscess due to Corynebacterium equi. Report of the first case in a patient with acquired immune deficiency syndrome, Am. J. Med., 80, 685–688, 1986. 113. Ellenberger, M.A., Kaeberle, M.L., and Roth, J.A., Equine cell-mediated immune response to Rhodococcus (Corynebacterium) equi, Am. J. Vet. Res., 45, 2424–2427, 1984. 114. Aihara, H., Watanabe, K., and Nakamura, R., Characterization of production of cholesterol oxidases in three Rhodococcus strains, J. Appl. Bacteriol., 61, 269–274, 1986. 115. Arlotti, M. et al., Rhodococcus equi infection in HIV-positive subjects: A retrospective analysis of 24 cases, Scand. J. Infect. Dis., 28, 463–467, 1996. 116. Scotton, P.G. et al., Rhodococcus equi nosocomial meningitis cured by levofloxacin and shunt removal, Clin. Infect. Dis., 30, 223–224, 2000. 117. Phumoonna, T. et al., Chimeric vapA/groEL2 DNA vaccines enhance clearance of Rhodococcus equi in aerosol challenged C3H/He mice, Vaccine, 26, 2457–2465, 2008. 118. Takai, S. et al., Detection of virulent Rhodococcus equi in tracheal aspirate samples by polymerase chain reaction for rapid diagnosis of R. equi pneumonia in foals, Vet. Microbiol., 61, 59–69, 1998. 119. Monego, F. et al., Molecular characterization of Rhodococcus equi from horse-breeding farms by means of multiplex PCR for the vap gene family, Curr. Microbiol., 58, 399–403, 2009. 120. Staden, R., Beal, K.F., and Bonfield, J.K., The Staden package, 1998, Methods Mol. Biol., 132, 115–130, 2000. 121. Zheng, H., Tkachuk-Saad, O., and Prescott, J.F., Development of a Rhodococcus equi-Escherichia coli plasmid shuttle vector, Plasmid, 38, 180–187, 1997. 122. Mangan, M.W., Byrne, G.A., and Meijer, W.G., Versatile Rhodococcus equi–Escherichia coli shuttle vectors, Antonie van Leeuwenhoek, 87, 161–167, 2005. 123. Dabbs, E.R., Gowan, B., and Andersen, S.J., Nocardioform arsenic resistance plasmids and construction of Rhodococcus cloning vectors, Plasmid, 23, 242–247, 1990.

15 Saccharopolyspora Caroline Duchaine and Yvon Cormier CONTENTS 15.1 Introduction...................................................................................................................................................................... 169 15.1.1 Classification......................................................................................................................................................... 169 15.1.2 Clinical Features and Pathogenesis.......................................................................................................................170 15.1.3 Diagnosis...............................................................................................................................................................171 15.2 Methods.............................................................................................................................................................................171 15.2.1 Sample Preparation................................................................................................................................................171 15.2.2 Detection Procedures.............................................................................................................................................171 15.3 Conclusion........................................................................................................................................................................ 172 References.................................................................................................................................................................................. 172

15.1  INTRODUCTION 15.1.1  Classification In agricultural environments, thermophilic actinomycetes grow in heated moist hay, grain, and straw, and are thus pre­ sent in the bioaerosols of dairy barns during material handling. “Farmer’s lung,” one of the most common forms of hypersensitivity pneumonitis,1,2 is usually caused by Saccharopolyspora rectivirgula, a thermophilic actinomycete.2,3 S. rectivirgula has also been associated with a respiratory disease in horses known as heaves.4,5 Aerobiological and environmental studies are frequently performed to evaluate the presence of S. rectivirgula in the work environment of farmers and thus also to assess exposure.6–8 Identification of actinomycetes is difficult using traditional microbiological approaches since most of their differential characteristics are structural and identification requires fastidious techniques such as electron microscopy or slide culture. S. rectivirgula can be differentiated from the closest affiliated species by a combination of tests. Nowadays, molecular biology approaches, including sequencing of the 16S rRNA gene, allow for the identification of isolates. In 1987, gram-positive bacteria with high G + C content were grouped to form a new class (Actinomycetes) composed of three groups: Bifidobacteria, Propionibacteria, and Actinomycetales. The latter included several families, such as Pseudonocardiaceae, which shelters the genus Saccharopolyspora. The actinomycetes, which were included in the Actinomycetales family, were grouped for their ability to form branched hyphae during one of their division stage. S. rectivirgula mycelium, when mature, is fragmented into rods and cocci by septation and hyphae disarticulation. Actinomycetes harboring this characteristic are called “nocardioforms,” since species from the Nocardia genus exhibit the same characteristics.9

The genus Micropolyspora was created in 1961 to group actinomycetes producing short chains of spores in the aerial and substrate mycelium.10 Pepys and collaborators established the relationship between a thermophilic actinomycete and Farmer’s lung disease for the first time in 1963.1 Thermopolyspora polyspora was the first name given to this Farmer’s lung agent.11 This name was subsequently rejected because the species description was obtained using contaminated strains.12 Micropolyspora faeni was the proposed name for this thermophilic actinomycete (meaning: isolated from hay (“foin” in French) by Cross et al.13 During the same period, Russian scientists isolated thermophilic actinomycetes from soil samples.14 These actinomycetes were placed in the Thermopolyspora genus following morphological analyses as well as cell wall chemical composition. The isolated strains named T. rectivirgula were later placed in the Micropolyspora genus,15 and, following new analyses, were named Micropolyspora rectivirgula. M. rectivirgula had never been implicated in any lung affection. In 1977, type species Micropolyspora brevicatena was reclassified because of the presence of mycolic acid in its cell wall.16 The loss of the type species in the genus Micropolyspora was followed by the dissolution of the genus and the overall reclassification of the other species, including the overall comparison and classification of Micropolyspora faeni and Micropolyspora rectivirgula. The two species showing high resemblance and similarities in their morphological features,17 phage susceptibility,18 comparative taxonomic and serologic studies were performed in 1979 and the two species were considered as synonymous.19 The International Code of Nomenclature of Bacteria indicates that the earliest name has priority when synonyms are discovered. However, even if the epithet rectivirgula had been published prior to the epithet faeni, the synonym species description was proposing 169

170

the epithet faeni priority: this name was thus proposed as nomen conservandum according to International Code of Nomenclature of Bacteria of 1975. Finally, at this time, even the presence of more than 200 peer-reviewed publications on Farmer’s lung disease and involving the species M. faeni did not weigh enough for the nomen conservandum to be accepted and the request was rejected. The earliest name was thus conserved and Micropolyspora rectivirgula was kept. Since the type species of the Micropolyspora genus (M. brevicatena) had just been reclassified in the Nocardia genus (and named Nocardia brevicatena), the genus was no more valid and M. rectivirgula had to be placed in a new genus and its name had to be changed again. A new genus was proposed based on this species (Faenia) and Faenia rectivirgula was proposed as the type species for the new name.20 The genus Faenia was described as an actinomycete producing branched aerial and substrate mycelium, 0.8–1.2 µm diameter, with spore chains (2–15) on both mycelia, gram-positive, and growing from 30°C to 60°C. The cell wall is of type IV: meso- diaminopimelic acid (meso-DAP), galactose, arabinose, and no nocardiomycolic acids. F. rectivirgula (type species) was described in 1983. In addition to the genus characteristics, it has smooth, ovoid, or spherical spores that are 0.7–1.5 µm diameter and form straight chains (never spirals). Older hyphae and spores may exhibit a negative response to Gram staining. Colonies are yellow-brown to pale orange and aerial mycelium is white. Growth is possible between 33°C and 60°C and optimal growth temperature is between 50°C and 55°C. In 1984, Goodfellow established a classification of actinomycetes and grouped all the genera containing mesoDAP, arabinose and galactose, and lacking mycolic acids in a group named micropolyspora (unofficial name).21 In 1988, Embley et al.22 revised this unofficial group and created the Pseudonocardiaceae family. It now includes Actinopolyspora, Amylocolatopsis, Faenia, Pseudonocar­ dia, Saccharomonospora, and Saccharopolyspora. This family exhibits all the characteristics proposed in 1984. Actinomycetes phylogeny was established using the 16S rRNA gene analyses.22,23 The same authors24 established that Faenia rectivirgula, Saccharopolyspora hirsute, and Pseudonocardia thermophila formed a very tight phylogenetic group of actinomycetes, and possessed more affinity between each other than with the other members of Pseudonocardiaceae family. Chemotaxonomic analyses also demonstrated high similitude between Saccharopolyspora and Faenia.25 In 1989, Korn-Wendish et al. performed a comprehensive comparison of Faenia rectivirgula with species of the genus Saccharopolyspora (S. taberi, S. erythrea, and S. hirsuta), as well as with Pseudonocardia thermophila, Saccharomonospora viridis, and Amylocalatopsis azurea. This study included morphological, physiological, biochemical (DAP), ADN G + C content, phage sensitivity, enzyme and protein profiles, and restriction enzyme digestion polymorphism data demonstrated that F. rectivirgula showed enough similarities with members of the genus Saccharopolyspora

Molecular Detection of Human Bacterial Pathogens

to be transferred to it.26 The Saccharopolyspora rectivirgula species description was finally published; its main features are that it is aerobic, gram-positive, nonacid-fast, with a type IV cell wall that includes meso-DAP, arabinose and galactose, and no mycolic acid. Substrate mycelium is yellow to orange on tryptic soy agar and is well developed, branched, and septed with a diameter from 0.5 to 0.8 µm. Aerial mycelium is white to pale pink and develops in the presence of 0.5% NaCl. Spores are present on both mycelium types and are in chains on short, unbranched, lateral or terminal sporophores. Spore surface is rough. Strains of S. rectivirgula are thermophilic and grow at temperatures between 37°C and 63°C, with optimum growth at 50°C. SR grows in the presence of 10% NaCl and is lysozyme sensitive. Esculin, arbutin, and urea are hydrolyzed. The G + C content is 70.4 mol%. Recently, Yassin identified a novel Saccharopolyspora specie (i.e., Saccharopolyspora rosea) from a human patient with bronchial carcinoma.27 The bacterium demonstrates morphological and chemotaxonomic characteristics consistent with those of members of the genus Saccharopolyspora, but is clearly distinguishable from the recognized members of the genus Saccharopolyspora by various biochemical tests. Comparative analysis of the 16S rRNA gene sequence revels that the type strain IMMIB L-1070(T) (=DSM 45226(T) = CCUG 56401(T)) displays 93.5%–96.9% sequence similarity with respect to members of the genus Saccharopolyspora, thus confirming its distinct phyletic position.27

15.1.2  Clinical Features and Pathogenesis Although S. rectivirgula is not a pathogenic bacterium per se, it is involved in an immunologic disease called hypersensitivity pneumonitis (HP), or extrinsic allergic alveolitis. HP is a very complex affection with clinical signs and symptoms varying in intensity and induced by a wide range of inhaled organic antigen. Its classical clinical presentation includes three major forms: acute, subacute, and chronic. Inhaled antigens can be derived from various sources such as hay, straw, bedding, water reservoirs, and animal dander or proteins. The most common forms of HP are “bird fancier’s disease” (induced by bird antigens) and farmer’s lung disease (induced by S. rectivirgula). HP is recognized by different clinical features. In the acute form, patients develop cough, shortness of breath (dyspnea), fever, and chills 3–8 h after exposure to the causative antigen. The subacute form is more insidious, usually lacking the fever and chills, and evolves into progressive dyspnea over weeks to months. The chronic form is usually reserved for permanent sequela of long-term ongoing subacute disease or repeated bouts of acute HP. The pathophysiology of HP is still complex and incompletely understood. Basically, HP is a delayed cellular immune response characterized by the accumulation of activated lymphocytes in the lungs.28 The pathology is similar to nonspecific interstitial pneumonitis (NSIP), with the presence of ill-formed granulomas. In more chronic forms lung fibrosis29 and/or emphysemateous destruction can occur.30 The major HP treatment is still antigen contact avoidance by environmental quality

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control. In severe acute forms, oral corticosteroids will accelerate the initial recovery but corticosteroids probably do not alter the long-term outcome.31

15.1.3  Diagnosis Validated diagnostic criteria were recently published by Lacasse et al.32 These criteria are based on clinical presentation, environmental exposure, and specific serum antibodies to the suspected antigen. Bronchoalveaolar lavage, which yields large numbers of lymphocytes28 and typical interstitial patterns on high-resolution computed tomography,33 is usually done to confirm the clinical diagnosis. The isolation of S. rectivirgula from a patient with farmer’s lung has never been successful since this actinomycete is thermophilic and does not invade human lung tissues or grow in the lungs. Bronchoalveolar lavage will show inflammatory cells but when cultured, S. rectivirgula is not recovered. However, a recent report shows the detection of b-glucans, a mould surface antigen, in the BAL of a patient with HP due to moulds exposure.34 The causative agent can be searched for in the work environment of a patient suffering farmer’s lung through air or bulk samples (hay, straw, bedding material, compost, etc.). The search for the SR is however usually not required; its presence and exposure are indirectly confirmed by the identification of specific IgG antibodies in the serum of the exposed subject. To confirm exposure to S. rectivirgula infection, air or bulk samples are often obtained from the dairy barn or animal house where patient has been exposed. The samples are plated on Tryptic soy agar and incubated at 52°C for 5 days. Colonies of S. rectivirgula are recognized by their yellowish/brown color and white aerial mycelium. Biochemical identification of S. rectivirgula relies on the metabolic reactions, cell membrane lipids, and colony and cell morphology. Cellular fatty acids (CFA) are structural components present in the membrane or cell wall. When cells are cultured in standard conditions, lipid composition is reproducible at the genus, species, or subspecies levels. CFA are analyzed using gas chromatography and comparison databases are available (MIDI Labs).35 Soluble proteins profile analysis has been described.36 Briefly, polyacrylamide gel electrophoresis in denaturing conditions of a protein suspension obtained after cell breaking using glass beads or ultrasonication is used for profile comparison purposes. This type of analysis may lead to numerical analysis37 and the determination of dendrograms and similarity profiles. Biochemical tests were validated.38 These authors proposed an identification algorithm using colony and cell morphologies, macromolecule degradation, utilization of sugars, acid production from the sugars, and antibiotic resistance. Those tests allow the differentiation between the agents most frequently responsible for HP cases Saccharopolyspora, Thermoactinomyces, and Saccharomonospora. Serological techniques such as electrosyneresis (ES), ouchterlony double diffusion (DD), ELISA, and Western blot (WB) have also been employed for

identification of Saccharopolyspora rectivirgula associated with farmer’s lung disease.39 Due to the limited value and the lack of reproducibility of some classical biochemical tests, Duchaine et al.40 proposed a simple and rapid scheme for the identification of S. rectivirgula: optimal growth temperature (55°C); colony appearance based on morphology (filamentous) and color (beige to orange-brown); microscopic morphology (chains of spores on both aerial and substrate mycelium); growth on NaCl 10%; cell wall analysis (type IV); and the verification of antibody response with serum from a patient with farmer’s lung. This last criterion is important to confirm the immunogenicity of the strains identified as S. rectivirgula. This scheme provides an accurate and efficient way of identifying S. rectivirgula strains and evaluating exposure to this bacterium. When Faenia rectivirgula was transferred into the Saccharopolyspora genus, Korn-Wendisch et al.26 published a complete description of S. rectivirgula in comparison with the other species regarding various markers (morphology, biochemistry, DNA, physiology, phage susceptibility patterns, enzyme and protein patterns, plasmids, and restriction fragments length polymorphisms). Given the difficulty and time consuming nature of differentiating members of the family Pseudonocardiaceae on the basis of micromorphology and biochemical properties, molecular techniques have been increasingly applied. Morón et al.41 reported the use of genus-specific primers from 16S rRNA gene for PCR identification of Saccharopolyspora. Random amplified ribosomal DNA restriction analysis approach has been published42 in the differentiation of closely related thermophilic actinomycetes involved in HP disease but has not been used in any other published report. Although this information is available, some authors used to base their identification of S. rectivirgula on colony morphology.43 Amplified ribosomal DNA restriction analysis (ARDRA) is also valuable for identification of environmental actinomycete-like organisms, including Saccharopolyspora rectivirgula and Saccharomonospora viridis.43

15.2  METHODS 15.2.1  Sample Preparation Air or bulk samples are obtained from dairy barn or animal house where the patient has been exposed to farmer’s lung. Several air sampling approaches have been described.7

15.2.2  Detection Procedures (i) Genus-Specific PCR  Morón et al.41 employed genus-specific primers for PCR identification of Saccharopolyspora. The primers are derived from the conserved region of the 16S rRNA gene—that is, forward primers SP1 (5′-GTGGAACCCATCCCCACACC-3′) and SP2 (5′-GTGGAAMCAGTCCCCACACC-3′)

172

both at positions 801–819, and reverse primer SP3 (5′-GGTGACGGTAGGTGTAGAAG-3′) at positions 450–469. The expected size of the PCR product using either SP1/SP3 or SP2/SP3 as amplification primers is 371 bp. Procedure 1. PCR mixture (25 µL) is made up of 0.2 mM each of the four dNTPs, 0.1 µM each primer, 5  µL extracted DNA, and 0.5 U Taq polymerase (Appligene) with the appropriate reaction buffer. Controls without bacterial DNA are included for each PCR run. 2. Amplification is conducted in a Perkin Elmer Cetus DNA Thermal Cycler 480 with 40 cycles of 93°C for 30 s, 53°C for 30 s, and 72°C for 2 min, followed by 10 min at 72°C. 3. The resulting amplicons are analyzed by electrophoresis (0.25 V/cm) in 1.2% (w/v) agarose gels, and stained with ethidium bromide. Note: The product of 371 bp is obtained with both pairs of primers, SPl/SP3 and SP2/SP3, from all the Saccharo­ polyspora  strains tested, including Saccharopolyspora erythraea, Saccharopolyspora gregorii, Saccharopolyspora hirsuta subsp. hirsuta, Saccharopolyspora hirsuta kobensis, Saccharopolyspora hordei, Saccharopolyspora rectivirgula, Saccharopolyspora spinosa, and Saccharopolyspora taberil. Two strains of Saccharopolyspora rectivirgula examined contain sequences around positions 450–469 that do not perfectly match with SP3 (nucleotide positions 464 and 465), but still produce the 371 bp amplification fragment. The effect of this divergence is only observed when the annealing temperature is increased by at least 4°C. In this case, experiments done at annealing temperatures of 57°C or higher fail to produce amplification fragments. (ii) 16S rRNA Gene Sequence Analysis  Paster et  al.44 reported a pair of universal primers, 27F (5′-AGAGTTTG ATCCTGGCTCAG-3′, positions 8 to 27 in Escherichia coli 16S rRNA gene) and 1492R (5′-TACCTTGTTACGACTT-3′, positions 1492 to 1507), which cover nearly the entire length of the gene the 16S rRNA from bacteria. Sequencing analysis of the resulting amplicon facilitates the identification of bacteria including Saccharopolyspora spp. Alternative 16S rRNA gene primers described by Frank et al.45 may be also utilized. Procedure 1. PCR mixture (100 µL) is made up of 50 mM KCl, 10 mM Tris pH 8.3, 1.5 mM MgCl2, 2 mM each of dNTP, 1 µM (each) primers 37F and 1492R, 2.5 U of Taq DNA polymerase, and 1 pg to 100 ng of template DNA. 2. The mixture is subjected to an initial denaturation at 94°C for 5 min, 25 cycles of 94°C for 60 s, 50°C for 30 s, and 72°C for 2 min, and a final extension step at 72°C for 5 min.

Molecular Detection of Human Bacterial Pathogens



3. The resulting PCR products are then sequenced by use of the ABI technology with the same universal primers. The nucleotide sequence is analyzed with BLAST program.

15.3  CONCLUSION Saccharopolyspora rectivirgula (Micropolyspora faeni) is one of the major agents responsible for farmer’s lung disease, a form of hypersensitivity pneumonitis that results from repeated inhalation of antigens from mouldy hay or straw. Although S. rectivirgula is capable of inducing a complex immunologic reaction in patients with farmer’s lung disease, and is detectable in the surrounding environment of patients, its precise role in the pathogenesis of farmer’s lung disease is still unclear. Due to its variable phenotypic characteristics, identification of Saccharopolyspora strains on the basis of classical tests is difficult and often inconclusive. In particular, microscopic and macroscopic examinations require specialized skills and the assessment of growth properties; macromolecule degradation, citrate utilization, and acid production from carbohydrates are time consuming and expensive. Although fatty acid analysis is very useful for the identification of Saccharopolyspora strains, it is not readily available outside of well-resourced research or reference laboratories. In recent years, molecular techniques have been applied for rapid and specific detection of Saccharopolyspora bacteria. Future development of real-time PCR will not only allow convenient detection and quantitation of Saccharopolyspora bacteria, but also reveal new insights on the role of these bacteria in farmer’s lung disease.

REFERENCES



1. Pepys, J. et al., Farmer’s lung: Thermophilic actinomycetes as a source of “farmer’s lung hay” antigen, Lancet, 1, 12, 1963. 2. Lacasse, Y., and Cormier, Y., Hypersensitivity pneumonitis, Orphanet J. Rare Dis., 1, 25, 2006. 3. Lacasse, Y. et al., Classification of hypersensitivity pneumonitis: A hypothesis, Int. Arch. Allergy Immunol., 149, 161, 2009. 4. Couetil, L.L. et al., Inflammatory airway disease of horses, J. Vet. Intern. Med., 21, 356, 2007. 5. Derksen, F.J. et al., Airway reactivity in ponies with recurrent airway obstruction (heaves), J. Appl. Physiol., 58, 598, 1985. 6. Cormier, Y. et al., Airborne microbial contents in two types of swine confinement buildings in Quebec, Am. Ind. Hyg. Assoc. J., 51, 304, 1990. 7. Duchaine, C., Grimard, Y., and Cormier, Y., Influence of building maintenance, environmental factors, and seasons on airborne contaminants of swine confinement buildings, Aihaj, 61, 56, 2000. 8. Kang, Y.-J., and Frank, J.F., Biological aerosols: A review of airborne contamination and its measurement in dairy processing plants, J. Food Prot., 52, 512, 1989. 9. Prauser, H., Nocardioides, a new genus of order Actinomycetales, Int. J. Syst. Bacteriol., 26, 58, 1976. 10. Lechevalier, H.A., Solotorovsky, M., and McDurmont, C.L., A new genus of actinomycetales: Micropolyspora gen. nov., J. Gen. Microbiol., 26, 11, 1961.

Saccharopolyspora 11. Corbaz, R., Gregory, P.H., and Lacey, M.E., Thermophilic and mesophilic Actinomycetes in mouldy hay, J. Gen. Microbiol., 32, 449, 1963. 12. Henssen, A., Importance of thermophilic microorganisms for the decomposition of stable manure, Arch. Mikrobiol., 27, 63, 1957. 13. Cross, T., Maciver, A.M., and Lacey, J., The thermophilic actinomycetes in mouldy hay: Micropolyspora faeni sp. nov., J. Gen. Microbiol., 50, 351, 1968. 14. Krassilnikov, N.A., and Agre, N.S., On two new species of Thermopolyspora, Hindustan Antibiotics Bull., 6, 97, 1964. 15. Lechevalier, H.A., and Lechevalier, M.P., Biology of actinomycetes, Ann. Rev. Microbiol., 21, 71, 1967. 16. Collins, M.D. et al., Distribution of menaquinones in actinomycetes and corynebacteria, J. Gen. Microbiol., 100, 221, 1977. 17. Dorokhova, L.A. et al., Morphology of 2 cultures belonging to the Micropolyspora genus, Mikrobiologiia, 39, 95, 1970. 18. Prauser, H., and Momirova, S., Phage sensitivity, cell wall composition and taxonomy of various thermophilic actinomycetes, Zeitschrift Allgemeine Mikrobiol., 10, 219, 1970. 19. Jones, M.P., McCarthy, A.J., and Cross, T. Taxonomic and serologic studies on Micropolyspora faeni and Micropolyspora strains from soil bearing the specific epithet rectivirgula, J. Gen. Microbiol., 115, 343, 1979. 20. Kurup, V.P., and Agre, N.S., Transfer of Micropolyspora rectivirgula (Krassilnikov and Agre 1964) Lechevalier, Lechevalier, and Becker 1966 to Faenia gen. nov., Int. J. Syst. Bacteriol., 33, 663, 1983. 21. Goodfellow, M., and Cross, T. Classification. In Goodfellow, M., Mordarski, M., and Williams, S.T. (Editors), The Biology of Actinomycetes, pp. 7–164, Academic Press, London, 1984. ‌ 22. Embley, M.T., Smida, J., and Stackebrandt, E., The phylogeny of Mycolate-less wall chemotype IV Actinomycetes and description of Pseudonocardiaceae fam. nov., Syst. Appl. Microbiol., 11, 44, 1988. 23. Fox, G.E., and Stackebrandt, E., The application of 16 S rRNA cataloguing and 5 S rRNA sequencing in bacterial systematics, Methods Microbiol., 19, 405, 1987. 24. Embley, T. et al., Chemotaxonomy of wall type IV actinomycetes which lack mycolic acids, J. Gen. Microbiol., 134, 953, 1988. 25. Embley, T.M., Smida, J., and Stackebrandt, E., Reverse transcriptase sequencing of 16S ribosomal RNA from Faenia rectivirgula, Pseudonocardia thermophila and Saccharopolyspora hirsuta, three wall type IV actinomycetes which lack mycolic acids, J. Gen. Microbiol., 134, 961, 1988. 26. Korn-Wendish, F. et al., Transfer of Faenia rectivirgula Kurup and Agre 1983 to the genus Saccharopolyspora Lacey and Goodfellow 1975, elevation of Saccharopolyspora hirsuta subso. taberi Labeda 1987 to species level, and emended description of the genus Saccharopolyspora, Int. J. Syst. Bacteriol., 39, 430, 1989. 27. Yassin, A.F., Saccharopolyspora rosea sp. nov., isolated from a patient with bronchial carcinoma, Int. J. Syst. Evol. Microbiol., 59, 1148, 2009. 28. Cormier, Y. et al., Bronchoalveolar lavage in farmers’ lung disease: diagnostic and physiological significance, Br. J. Ind. Med., 43, 401, 1986.

173 29. Perez-Padilla, R. et al., Mortality in Mexican patients with chronic pigeon breeder’s lung compared with those with usual interstitial pneumonia, Am. Rev. Respir. Dis., 148, 49, 1993. 30. Lalancette, M. et al., Farmer’s lung. Long-term outcome and lack of predictive value of bronchoalveolar lavage fibrosing factors, Am. Rev. Respir. Dis.,148, 216, 1993. 31. Monkare, S., Influence of corticosteroid treatment on the course of farmer’s lung, Eur. J. Respir. Dis., 64, 283, 1983. 32. Lacasse, Y. et al., Clinical diagnosis of hypersensitivity pneumonitis, Am. J. Respir. Crit. Care Med., 168, 952, 2003. 33. Lynch, D.A. et al., Hypersensitivity pneumonitis: Sensitivity of high-resolution CT in a population-based study, Ajr, 159, 469, 1992. 34. Ashitani, J. et al., Elevated levels of beta-D-glucan in bronchoalveolar lavage fluid in patients with farmer’s lung in Miyazaki, Japan, Respiration; Int. Rre. Tthoracic Dis., 75, 182, 2008. 35. Bernard, K.A., Bellefeuille, M., and Ewan, E.P., Cellular fatty acid composition as an adjunct to the identification of asporogenous, aerobic gram-positive rods, J. Clin. Microbiol., 29, 83, 1991. 36. Moore, W.E. et al., Polyacrylamide slab gel electrophoresis of soluble proteins for studies of bacterial floras, Appl. Environ. Microbiol., 39, 900, 1980. 37. Kersters, K., and De Ley, J., Identification and grouping of bacteria by numerical analysis of their electrophoretic protein patterns, J. Gen. Microbiol., 87, 333, 1975. 38. Kurup, V.P., and Fink, J.N., A scheme for the identification of thermophilic actinomycetes associated with hypersensitivity pneumonitis, J. Clin. Microbiol., 2, 55, 1975. 39. Reboux, G. et al., Assessment of four serological techniques in the immunological diagnosis of farmers’ lung disease, J. Med. Microbiol., 56, 1317, 2007. 40. Duchaine, C. et al., Saccharopolyspora rectivirgula from Quebec dairy barns: Application of simplified criteria for the identification of an agent responsible for farmer’s lung disease, J. Med. Microbiol., 48, 173, 1999. 41. Morón, R., González, I., and Genilloud, O., New genus-specific primers for the PCR identification of members of the genera Pseudonocardia and Saccharopolyspora, Int. J. Syst. Bacteriol., 49, 149, 1999. 42. Harvey, I. et al., Random amplified ribosomal DNA restriction analysis for rapid identification of thermophilic Actinomycetelike bacteria involved in hypersensitivity pneumonitis, Sys. Appl. Microbiol., 24, 277, 2001. 43. Kurtboke, D.I., Murphy, N.E., and Sivasithamparam, K., Use of bacteriophage for the selective isolation of thermophilic actinomycetes from composted eucalyptus bark, Can. J. Microbiol., 39, 46, 1993. 44. Paster, B.J. et al., Prevalent bacterial species and novel phylotypes in advanced noma lesions, J. Clin. Microbiol., 40, 2187, 2002. 45. Frank, J.A. et al., Critical evaluation of two primers commonly used for amplification of bacterial 16S rRNA genes, Appl. Environ. Microbiol., 74, 2461, 2008.

16 Streptomyces Angel Manteca, Ana Isabel Pelaez, Maria del Mar Garcia-Suarez, and Francisco J. Mendez CONTENTS 16.1 Introduction.......................................................................................................................................................................175 16.1.1 Classification, Morphology, Biology, and Epidemiology......................................................................................175 16.1.2 Clinical Features and Pathogenesis.......................................................................................................................175 16.1.3 Diagnosis.............................................................................................................................................................. 177 16.2 Methods.............................................................................................................................................................................178 16.2.1 Sample Preparation................................................................................................................................................178 16.2.2 Detection Procedures.............................................................................................................................................178 16.3 Conclusion and Future Perspectives................................................................................................................................. 179 Acknowledgments...................................................................................................................................................................... 179 References.................................................................................................................................................................................. 179

16.1  INTRODUCTION 16.1.1  C  lassification, Morphology, Biology, and Epidemiology Actinomycetales are gram-positive, mycelium-forming, soil bacteria that play an important role in mineralization processes in nature. Streptomyces is an extremely important bacterium in biotechnology because approximately twothirds of industrial antibiotics are synthesized by members of this genus.1 Streptomycetes present a complex development cycle that includes programmed cell-death processes, as well as differentiation phenomena that lead to sporulation2–4 (Figure 16.1). After spore germination, a fully compartmentalized mycelium starts to grow and early on suffers an orderly death process in which live and dead segments alternate in the same hyphae.2–4 Subsequently, the viable segments start to enlarge as a multinucleated mycelium that grows in successive waves that determine the characteristic complex growth curves of this microorganism. Two stages of secondary multinucleated mycelium have been defined: early secondary mycelium that does not yet have the hydrophobic layers characteristic of aerial hyphae and late secondary mycelium that starts to express the genes encoding for the chaplin and rodlin proteins, which are mainly responsible for the formation of the hydrophobic layer characteristic of aerial structures.5 The traditionally denominated substratevegetative mycelium corresponds to the early secondary multinucleated mycelium, which still lacking the hydrophobic layers characteristic of the aerial mycelium and would match therefore to the late secondary mycelium which has acquired those hydrophobic covers.6 A vegetative role has

been postulated to these compartmentalized hyphae by contrast to the secondary multinucleated mycelium that should be considered the reproductive structure, since it is destined to sporulate.7 The secondary mycelium has been described also as the antibiotic producing.7 Streptomyces displays an unequivocally morphology characterized by the formation of branched, thin hyphae of 0.5–2 µm in diameter, and spore chains. Isolated colonies of this bacterium are also easy to identify as they exhibited grayish-white appearance, and a medium size around 3 mm in diameter (Figure 16.2a). Streptomycetes are infrequently encountered in clinical practice but are important potential causes of serious human and animal infections.10,11 Their natural habitats are mainly natural soils, and they are universally distributed in environment. The ubiquitous nature and the low pathogenicity of Streptomyces organisms make most clinical isolates contaminants or colonizers. Despite that, at least in our experience, Streptomyces strains are often not recognized in hospitals, and several Streptomyces infections and colonizations are not being identified. In the Oviedo Central Hospital (Spain), we have described three such clinical cases during last 5 years (2004–2009)8,9,12; in the Section of Clinical Microbiology, University of Texas, six cases have been described during 9  years (1997–2005)11; and probably many more cases are either not being identified or they are considered as contaminants.

16.1.2  Clinical Features and Pathogenesis As stated earlier, Streptomycetes are rarely documented as cause of human disease. The most common manifestation 175

176

Molecular Detection of Human Bacterial Pathogens

MI MII with hydrophobic covers

MII

SUBSTRATE MI

MII Multinucleated second mycelium

Compartmentalized first mycelium First death round

AERIAL MII Multinucleated second mycelium with hydrophobic covers

Spores Spores with hydrophobic covers

Second death round

FIGURE 16.1  Cell-cycle features of the Streptomyces development. Mycelial structures (MI, first compartmentalized mycelium; MII, second multinucleated mycelium), cell-death processes, spores, and traditional denomination (“substrate” and aerial) are indicated. (a)

(b)

FIGURE 16.2  Streptomyces morphology. (a) Colonies of Streptomyces cinereoruber isolated from a patient with severe chronic obstructive pulmonary disease.8 (b) Laser confocal observations of silicone prothesis infected with Streptomyces spp.9 Samples are stained with the viability assay LIVE/DEAD Bac-Light Bacterial Viability kit (Molecular Probes) (left), or in differential interference contrast mode (right). Arrows illustrate the border of the silicone biopsy.

of Actinobacteria infections are actinomycetomas which are generally easy to identify by their clinical presentation: chronic subcutaneous inflammatory tumor with discharging sinuses. The commonest affected sites are the foot, knee, thigh, hand, and arm. Lung-associated actinomycetoma are extremely rare and had been attributed to members of the Nocardia genus,13 and some Streptomyces species, such as Streptomyces pneumoniae or Streptomyces cinereoruber.8,14 Recently, a silicone mammary implant actinomycetoma has been also reported to Streptomyces.9 Most of these cases had

in common some degree of immunosuppression. In the last 20 years, at least 10 species belonging to the Streptomyces genus have been isolated from mycetomic and nonmycetomic infections.14,15 S. somaliensis is a well-documented agent in actinomycetomas.8,16 S. albus has been also cited as a human pathogen and the recent literature includes cases of extrinsic alveolitis and skin hypersensitivity type II in farm workers.12 Pulmonary infections due to Streptomyces in nonimmunocompromised hosts have been also reported9,14,17,18 (Table 16.1).

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Streptomyces

TABLE 16.1 Examples of Well-Documented Streptomyces Infections Isolate S. albus S. cinereoruber Streptomyces spp. S. maritimus S. albus S. nobilis S. humidus S. indigoferus Streptomyces spp. S. penumonia S. coelicolor

Type of Infection

Immunosuppression and/or Antineoplastic Treatments

Reference

Actinomycetoma forearm Lung coinfection Silicone mammary implant infection Lung nodule Lung nodule CVC-related boodstream infection CVC-related boodstream infection CVC-related boodstream infection Hypersensitivity pneumonitis Lung infection Brain abscess

Yes No Yes Yes Yes Yes Yes Yes Yes No NR

12 8 9 11 11 11 11 11 11 14 19

Abbreviations:  CVC, central venous catheter; NR, non reported.

Streptomycetes are most commonly limited to causing actinomycotic mycetoma in areas of the body that come into contact with soil, or those traumatized gain access to subcutaneous tissues through abrasions in the skin. Initial formation of painless nodules is followed by fluctuance and progression to formation of draining sinuses. The sinuses begin to drain, with the exudates often containing hard, compacted, filamentous microcolonies or granules. Additional sinus tracts and nodules may form, and the involved areas eventually become deformed by fibrous tissue. The process remains characteristically pyogenic rather than granulomatous and may reach deeper connective tissue, muscle, and bone. The disease rarely spreads hematogenously via the lymphatics.11 The treatment of Streptomyces infections remains a problem; there are no approved breakpoint antibiotics for Streptomyces species, and the correlation between the data obtained in vitro and the clinical evolution remains unclear.10 Many drugs are being used with variable results. The drugs used most frequently include trimethoprim-sulfamethoxazole (TS), dapsone, amikacin alone or combined with TS, and amoxicillin-clavulanic acid.15 All treatments have in common the need for long courses of therapy to obtain satisfactory results (Table 16.2). Our experience with three clinical cases is that prolonged treatment with TS (several weeks) is very effective (Table 16.2).8,9,12

16.1.3  Diagnosis Conventional Techniques. The first approach to diagnos­ ticate a Streptomyces infection/colonization is a direct examination of a biopsy or drainage material collected. Usually, granules (averaging 1 mm in diameter) are observed. A microscopic analysis can show gram-positive fine filaments.8,9,12 One of the main differences with fungal hyphae is that Streptomyces hyphae are much finer, having the diameter of a bacterium (0.5–2 µm) (Figure 16.2b). However, Streptomyces hyphae are sometimes very fragmented, and mycelia appearance is not evident.12

Streptomycetes are bacteria that are easily cultivable almost in all microbial culture media. However, and in order to facilitate the full differentiation and sporulation that facilitate colonies differentiation, poor media are preferred. Incubation temperatures of 28°C–30°C are recommended, and long incubation periods are necessary (around 1 week). In this sense, a good, easy-to-prepare media is GAE.8 Alternative culture media for Streptomyces cultures are detailed in Kieser et al. (Reference 20; see Methods.) Biochemical markers of taxonomic relevance in Streptomyces are, for example, utilization of cysteine, valine, phenylalanine, histidine, hydroxyproline, sucrose, mannitol, raffinose, dextran, or xylitol.1 Other activities with taxonomy relevance are lecithinase activity, lipolysis, pectin hydrolysis, nitrate reduction, elastin degradation, and more.1 Molecular Techniques. Identification of Streptomyces genus by the conventional techniques described in the preceding material is relatively easy. However, morphological, physiological, and biochemical approaches are not sufficient to TABLE 16.2 Antibiotic Susceptibility Data for the Cases Indicated in Table 16.1. The Number of Sensible Strains (11 in Total) and the Percentage for Each Antibiotic Are Indicated Antibiotic Imipenem Trimethroprim-sulfamethoxazole Erythromycin Clarithromycin Amikacin Amoxicilin-clavulanate Ceftriaxone Ampicilin Vancomycin Minocycline Tetracycline Vancomycin

Susceptibility 8 (73%) 6 (54%) 5 (45%) 4 (36%) 5 (45%) 4 (36%) 3 (27%) 3 (27%) 3 (27%) 2 (18%) 2 (18%) 2 (18%)

178

characterize strains and species in this genus, which is characterized by a broad biochemical and genetic diversity that traditionally has attributed to the description of more species than really have been validated by molecular techniques.20 For all these reasons, molecular techniques are beginning to be especially important for definitive identification and taxonomic characterization of Streptomyces strains. Also, molecular techniques are much faster, which can be also a critical advantage in the identification of this slow-growing bacterium. A first molecular approach to Streptomyces identification can be the analysis of amplification restriction of the 65-kDa fragment coding for the HSP.21 This kind of analysis can differentiate Streptomyces of other Actinomycetes and also works to identify some Streptomyces species, such as S. albus, S. griseus, and S. somaliensis.21 The fastest and most reliable methods in bacterial taxonomy and identification, and for identification of Streptomyces in particular, are those based in the analysis of 16S ribosomal DNA sequence: sequencing of 16S ribosomal RNA (rRNA)22 and analysis of the α-region of the 16S rRNA in the case of Streptomycetes (positions 158 to 276).23 The rRNA genes are essential for the survival of all organisms and are highly conserved in the bacterial and other kingdoms. Consequently, characterization of the 16s rRNA gene is now well established as a standard method for the identification of species, genera, and families of bacteria.24 This is reflected in the large body of information that has been accumulated on the phylogenetic analysis of microorganisms using this gene.25 In consequence, the full sequence of 16S rRNA can be considered as the master molecule for bacterial taxonomy.25 A rapid identification of 16S ribosomal DNA can be performed using the primers pA (5′-AGAGTTTGATCCTGGCTCAG-3′) and pH (5′-AAGGAGGTGATCCAGCCGCA-3′), as described previously.26 The α-region of the 16S rRNA has been described as one of the most variable in Streptomyces, and it is very useful for species identification.23 For this reason, when partial sequences of 16S rRNA are used for identification of this bacterium, they should include at least the α-region. Usually, correct 16S rRNA sequence analysis needs the elaboration of phylogenetic trees, including type 16S rRNA sequences from databases. For this end, phylogenetic trees based on maximum likelihood methods are the most recommended. For bacterial identification, 16S rRNA sequencing is definitive. It is the easiest, fastest, and most reliable technique for Streptomyces genus identification. However, when identification at the species level is necessary, additional experiments are usually needed. For example, the HSP RFLP indicated above, morphological and biochemical markers; RFLP from total DNA; DNA: DNA hybridizations; analysis of membrane fatty acids; enzyme patterns; and more.27

16.2  METHODS 16.2.1  Sample Preparation Streptomyces Cultivation and Isolation. The microorganisms should be isolated from the biological sample by

Molecular Detection of Human Bacterial Pathogens

growing them in solid media in which they have an entire life cycle with abundant sporulation, facilitating isolation and identification. We recommended poor media as GAE (glucose, asparagine, yeast extract)28 or SFM (Soya flour– mannitol).20 After agar addition, solid cultures are prepared in Petri dishes (8.5-cm diameter) as lawns on solid medium (30 mL/plate). Plates were incubated at 30°C at least during 7 days. Strains should be conserved as lyophilized spores.20 Microscopy. Isolated colonies or original sample (biopsy, sputum, exudates, etc.) can be observed directly at phase contrast microscope, or stained with Gram or other colorants. An alternative staining very useful in Streptomyces is the viability assay (LIVE/DEAD Bac-Light Bacterial Viability Kit, Molecular Probes, L-13152)2 (Figure 16.2b); a viability assay, based on staining cells in a population with propidium iodide and SYTO 9.2 Propidium iodide is specific for nucleic acids of damaged (leaky) cells, whereas SYTO 9 stains those of viable cells. The stain mixture is added directly to the samples on the slide. The coverslip is placed on top after staining for at least 10 min in the dark, and samples are observed under a fluorescence or confocal microscope at wavelengths of 488 and 568 nm for excitation, and 530 nm (green) or 630 nm (red) for emission. This fluorescence staining combined with confocal microscopy allows direct observation of Streptomyces hyphae in the original infected tissue, even if this tissue is very thick or dirty. An example of this kind of observation has been published recently for a Streptomyces infection in a silicone mammary implant.12 Physiological/Biochemical Phenotypes and Suscep­ tibility Tests. Physiological and biochemical phenotypes can be determined using standard bacterial identification techniques.1,29 Antimicrobial susceptibility testing can be performed simultaneously by Kirby Bauer disc diffusion30 or E-tests (AB Biodisk, Sweden). In the case of E-tests, a protocol for Streptomyces has been described by Martin et al.12

16.2.2  Detection Procedures Streptomyces DNA can be obtained from isolated colonies using standard methods.20 To obtain a rapid strain identification, PCR can be performed directly from a dilution of the isolated colonies, or even from a dilution of the original sample, biopsy, or wherever. The best primers for amplification of 16S rRNA are pA and pH.26 The 50  µL reaction mixture should contain 0.2 mM dNTPs mixture, 30 pmol/ µL of each primer (pA and pH); 1.5 mM MgCl2; 1x PCR buffer; 0.2 µL (1 U) Eco Taq DNA polymerase (Ecogen). The PCR program is as follows: denaturation at 95°C for 5 min; 30 cycles of 95°C for 1 min, 57°C for 1 min, and 72°C for 90 s; and final extension at 72°C for 10 min. The amplification products (1500 bp) is analyzed by horizontal gel electrophoresis in 0.8% agarose gel in Tris-Borate-EDTA (TBE), followed by ethidium bromide staining and visualization under UV light. PCR products from the extracted bands are purified by GFXTM PCR DNA and Gel Band Purification Kit (Amersham Biosciences). The amplicon is sequenced in

179

Streptomyces

an automatic sequencing as for example ABI PrismTM 3100 Genetic Analyzer (Perking Elmer). Sequence identification is done using the Gapped BLAST database search program facility of the National Center for Biotechnology Information (NCBI).31 For a more in-depth comparison between 16S rRNA sequences, phylogenetic trees are very useful. The most adequate phylograms are those based in nonparametric analysis [maximum likelihood methods (ML)]. An easy and robust free online platform for ML phylogenetic analyses can be found at http://phylemon. bioinfo.cipf.es/.32 Preliminary information can be obtained from isolates before sequencing. The 16S rRNA from different Streptomyces isolates can be compared by fragment length polymorphism (RFLP).33 Here, 16S rRNA are digested with restriction enzymes as HaeIII and AluI. An aliquot of each of the PCR-amplified 16S rRNA genes is digested at 37°C for 4 h, and the resulting restriction fragments separated on a 3% agarose electrophoresis gel. Identical patterns of bands are assigned to the same microorganism, whereas different patterns confirm those that are different Streptomyces species.

16.3  C  ONCLUSION AND FUTURE PERSPECTIVES The infections caused by Streptomyces species are rare and in general not very well documented. In this chapter, we described some of the most significant cases of Streptomyces infections as well as the methods for identification of these bacteria, with special emphasis on molecular techniques, which are especially important for the correct identification of this unusual pathogenic bacterium. Morphological, physiological, and biochemical approaches are not enough to characterize strains and species in this genus.20 The 16S rRNA sequencing and analysis is a culture-independent technique that can be performed directly from the primary material (i.e., biopsies, sputa, exudates). It is one of the most reliable and faster techniques for the correct identification of Streptomyces infections. Of all the most recent advances in molecular biology, 16S rRNA analysis is undoubtedly the technique of choice for identification of bacteria in general, and Streptomyces in particular, and in the near future, it will be increasingly implanted for bacterial clinical diagnosis. It should be noted that for identification at the species level, additional experiments are usually necessary, such as the HSP RFLP indicated previously; morphological and biochemical markers; RFLP from total DNA; DNA:DNA hybridizations; analysis of membrane fatty acids; or enzymatic patterns. Other innovative techniques that are being implanted in basic research will be progressively extended to clinical microbiological studies, including pathogenic Streptomyces strains. For instance, confocal microscopy, or Fourier transform infrared spectroscopy (FTIR) microscopy and even mass spectrometry techniques are becoming very important in the study of the biology of Streptomyces.2,3,34–36 These techniques will be very useful for the analysis of the clinical Streptomyces infections from direct biopsies, or other kind

of primary samples. FTIR can be also applied to species identification.34

ACKNOWLEDGMENTS This work was supported by grant MEC-06-BI0200615336-C04-02 and MEC-07-BIO2007-66313 from the DGI, Subdireccion General de Proyectos de Investigacion, Ministerio de Ciencia e Innovación, Spain. The authors would like to acknowledge Professor Jesus Sanchez (Area Microbiologia, Universidad de Oviedo) for the critical reading of the manuscript and help with phylogenetic analysis. A.M. was supported by a postdoctoral grant from Ministerio de Ciencia e Innovación, Spain; A.I.P. by grant CENIT07-CLEAM, Ministerio de Industria, Turismo y Comercio (CDTI), Spain; and M.M.G.S. was supported by grant MEC06-BIO2006–1533-C04–02.

REFERENCES













1. Locci, R. et al., Streptomycetes and related genera. In S.T. Williams, M.E. Sharpe and J.G. Holt (Editors), Bergey’s Manual of Systematic Bacteriology, Vol. 4, pp. 2451–2508, Williams & Wilkins Co., 1984. ‌ 2. Manteca, A., Fernandez, M., and Sanchez. J., A death round affecting a young compartmentalized mycelium precedes aerial mycelium dismantling in confluent surface cultures of Streptomyces antibioticus, Microbiology, 151, 3689, 2005. 3. Manteca, A., Fernandez, M., and Sanchez, J., Mycelium development in Streptomyces antibioticus ATCC11891 occurs in an orderly pattern which determines multiphase growth curves, BMC Microbiol., 5, 51, 2005. 4. Manteca, A. et al., A proteomic analysis of Streptomyces coelicolor programmed cell death, Proteomics, 6, 6008, 2006. 5. Manteca, A. et al., Aerial hyphae in surface cultures of Streptomyces lividans and Streptomyces coelicolor originate from viable segments surviving an early programmed cell death event, FEMS Microbiol. Lett., 274, 118, 2007. 6. Flärdh, K., and Buttner, M. J., Streptomyces morphogenetics: Dissecting differentiation in a filamentous bacterium, Nat. Rev. Microbiol., 7, 36, 2009. 7. Manteca, A., and Sanchez, J., Streptomyces development in colonies and soils, Appl. Environ. Microbiol., 75, 2920, 2009. 8. Manteca, A. et al., A rare case of lung coinfection by Streptomyces cinereoruber and Haemophilus influenzae in a patient with severe chronic obstructive pulmonary disease: Characterization at species level using molecular techniques, Diagn. Microbiol. Infect. Dis., 60, 307, 2008. 9. Manteca, A. et al., A rare case of silicone mammary implant infection by Streptomyces spp. in a patient with breast reconstruction after mastectomy: Taxonomic characterization using molecular techniques, Diagn. Microbiol. Infect. Dis., 63, 390, 2009. 10. McNeil, M.M., and Brown, J.M., The medically important aerobic actinomycetes: Epidemiology and microbiology, Clin. Microbiol, Rev., 7, 357, 1994. 11. Kapadia, M., Rolston, K.V., and Han, X.Y. Invasive Streptomyces infections: Six cases and literature review, Am. J. Clin. Pathol., 127, 619, 2007. 12. Martin, M.C. et al., Streptomyces albus isolated from a human actinomycetoma and characterized by molecular techniques, J. Clin. Microbiol., 42,5957, 2004.

180 13. Fuentes, A. et al., Report of five cases treated with imipenem or imipenem plus amikacin, Gac. Med. Mex., 142, 247, 2006. 14. Dunne, E.F., Burman, W.J., and Wilson, M.L., Streptomyces pneumonia in a patient with human immunodeficiency virus infection: Case report and review of the literature on invasive streptomyces infections, Clin. Infect. Dis., 27, 93, 1998. 15. Trujillo, M.E., and Goodfellow, M., Numerical phenetic classification of clinically significant aerobic sporoactinomycetes and related organisms, Antonie Leeuwenhoek, 84, 39, 2003. 16. Dieng, M.T. et al., Actinomycetomas in Senegal: Study of 90 cases, Bull Soc. Pathol. Exot., 98, 18, 2005. 17. Gugnani, A.C., Unaogu, I.C., and Emeruwa, C.N., Pulmonary infection due to Streptomyces griseus, J. Commun. Dis., 25, 38, 1993. 18. Nichols, W.G. et al., Fatal pulmonary infection associated with a novel organism, “para-streptomyces abscessus,” J.  Clin. Microbiol., 43, 5376, 2005. 19. Clarke, P.R. et al., Brain abcess due to Streptomyces griseus, J. Neurol. Neurosurg. Psychiatry, 27, 553, 1964. 20. Kieser, T. et al., Practical streptomyces genetics, Practical streptomyces genetics, pp. 2–5. The John Innes Foundation, Norwich, England, 2000. ‌ 21. Steingrube, V.A. et al., Rapid identification of clinically significant species and taxa of aerobic actinomycetes, including Actinomadura, Gordona, Nocardia, Rhodococcus, Streptomyces, and Tsukamurella isolates, by DNA amplification and restriction endonuclease analysis, J. Clin. Microbiol., 35, 817, 1997. 22. Edwards, U. et al., Isolation and direct complete nucleotide determination of entire genes. Characterization of a gene coding for 16S ribosomal DNA, Nucleic Acids. Res., 17, 7843, 1989. 23. Kataoka, M. et al., Application of the variable region in 16S rDNA to create an index for rapid species identification in the genus Streptomyces, FEMS Microbiol. Lett., 151, 249, 1997. 24. Amann, R.I., Ludwig, W., and Schleifer K.H., Phylogenetic identification and in situ detection of individual microbial cells without cultivation, Microbiol. Rev., 59, 143, 1995.

Molecular Detection of Human Bacterial Pathogens 25. Cole, J. R. et al., The Ribosomal Database Project: Improved alignments and new tools for rRNA analysis, Nucleic Acids Res., 37, 141, 2009. 26. Bruce, K.D. et al., Amplification of DNA from native populations of soil bacteria by using the polymerase chain reaction, App. Environ. Microbiol., 58, 3413, 1992. 27. Vandamme, P. et al., Polyphasic taxonomy, a consensus approach to bacterial systematic, Microbiol. Rev., 60, 407, 1996. 28. Mendez, C. et al., Role of substrate mycelium in colony development in Streptomyces, Can. J. Microbiol., 31, 446, 1985. 29. Murray, P. et al., Manual of Clinical Microbiology, 9th ed., American Society for Microbiology, Washington, DC, 2007. 30. Biemer, J.J., Antimicrobial susceptibility testing by the KirbyBauer disc diffusion method, Ann. Clin. Lab. Sci., 3, 135, 1973. 31. Zheng, Z. et al., A greedy algorithm for aligning DNA sequences, J. Comput. Biol., 7, 203, 2000. 32. Tarraga, J. et al., A suite of web tools for molecular evolution, phylogenetics and phylogenomics, Nucleic Acids. Res., 35, 38, 2007. 33. Broda, D.M., Musgrave, D.R. and Bell, R.G., Use of restriction fragment length polymorphism analysis to differentiate strains of psychrophilic and psychrotropic clostridia associated with blown pack’ spoilage of vacuum-packed meats, J. Appl. Microbiol., 88, 107, 2000. 34. Zhao, H. et al., Differentiation of Micromonospora isolates from a coastal sediment in Wales on the basis of Fourier transform infrared spectroscopy, 16S rRNA sequence analysis, and the amplified fragment length polymorphism technique, Appl. Environ. Microbiol., 70, 6619, 2004. 35. Huleihel, M., Pavlov, V., and Erukhimovitch, V., The use of FTIR microscopy for the evaluation of anti-bacterial agents activity, J. Photochem. Photobiol. B., 96, 17, 2009. 36. Freiwald, A., and Sauer, S. Phylogenetic classification and identification of bacteria by mass spectrometry, Nat. Protoc., 4, 732, 2009.

17 Tropheryma Shoo Peng Siah, Han Shiong Siah, and Dongyou Liu CONTENTS 17.1 Introduction.......................................................................................................................................................................181 17.1.1 Classification, Morphology, and Genome Structures............................................................................................181 17.1.2 Epidemiology........................................................................................................................................................ 182 17.1.3 Clinical Features................................................................................................................................................... 182 17.1.4 Diagnosis.............................................................................................................................................................. 183 17.1.4.1 Phenotypic Techniques.......................................................................................................................... 183 17.1.4.2 Genotypic Techniques............................................................................................................................ 183 17.2 Methods............................................................................................................................................................................ 184 17.2.1 Sample Preparation............................................................................................................................................... 184 17.2.2 Detection Procedures............................................................................................................................................ 184 17.3 Conclusion........................................................................................................................................................................ 187 References.................................................................................................................................................................................. 187

17.1  INTRODUCTION Whipple’s disease is a rare illness that occurs in less than one in 1,000,000 people and affects multiple organs and organ systems, predominantly in middle-aged men.1,2 It was first described postmortem by Dr. George H. Whipple in 1907 in a 36-year-old male, who died following a 5-year illness with fever, chronic cough, lymphadenopathy, and progressive weight loss.3 Up until 1947, Whipple’s disease was only diagnosed at autopsy, and in 1949, the bacterium was first detected as a period acid-Schiff positive (red staining) inclusion in foamy macrophages from an intestinal biopsy.4 Definitive confirmation of the pathogen was first obtained in 1961 by electron microscopic examination of the rod-shaped organism. Following the analysis of a short segment of the 16S ribosomal RNA (16S rRNA) gene by broad-range polymerase chain reaction (PCR) and sequencing, the identity of the bacterium was confirmed as a member of the actinomycetes, and the name Tropheryma whippeli was proposed in 1992.5 “Tropheryma” is derived from the Greek words for food (trophi) and barrier (eryma). The current name Tropheryma whipplei was adopted in 20016 after its successful culture and propagation in a human embryonic lung fibroblast cell line.7 Another group also reported the cultivation of the bacterium from cerebrospinal fluid.8 The completion of the whole genome sequences of two T. whipplei strains in 20039,10 rendered it possible to devise a cell-free culture system supplemented with amino acids for in vitro growth of the bacterium.11

17.1.1  Classification, Morphology, and Genome Structures Tropheryma whipplei was the only member of the genus Tropheryma, which is classified in the family Cellulomonadaceae, suborder Micrococcineae, order Actinomycetales, subclass Actinobacteridae, class Actino­ bacteria, phylum Actinobacteria. Sequence analysis of its 23S rRNA, 5S rRNA, DNAdependent RNA polymerase (rpoB), 65-kDa heat-shock protein (groEL2), elongation factor TU (tuf), ATPase beta subunit (atpD), and RNase P RNA (rnpb) genes placed Tropheryma whipplei phylogenetically among the high G + C content gram-positive bacterial class Actinobacteria.8,12,13 T. whipplei is a gram-positive bacterium although it is poorly stained due to its atypical envelope and thick cell wall.5 As such, it is commonly found to be gram-negative or gram-indeterminant. The bacterium has a unique ultrastructure unlike any other bacteria. Its membrane may be of bacterial or host origin as it lacks lipopolysaccharide, a feature that is shared by the macrophage plasma membrane.6,14 The bacterium measures 0.2–0.25 µm by 1.0–2.5 µm and occurs as periodic acid-Schiff positive inclusions in macrophages.15,16 Forming rope-like aggregates, the bacterium can be found intact extracellularly but are in various stages of degradation intracellularly.6 The bacterium multiplies extracellularly by perpendicular cell division but rarely within the macrophages.17 Electron microscopy showed that the extracellular bacteria in cell cultures adhere to each other via extracellular 181

182

matrix and tend to occur as aggregates. The bacterium is neither covered by flagella nor motile. Within infected cells, particularly in macrophages, it may be found in various stages of degradation within the vacuoles.6 The Tropheryma whipplei genome is the smallest within the Actinobacteria and DNA sequencing of the T. whipplei Twist and TW08/27 strains indicates that the circular genome is approximately 0.93 Mb in size and has a G + C content of approximately 46%, which is the lowest amongst the high G + C content gram-positive bacteria.9,10,18 The genome contains 808 predicted open reading frames. The pathway involved in synthesis of nine amino acids (histidine, tryptophan, leucine, arginine, proline, lysine, methionine, cysteine, and asparagine) are absent and synthesis of another seven (glutamate, glutamine, aspartate, threonine, valine, isoleucine, and phenylalanine) is partially deficient. The genes that encode components of the tricarboxylic acid cycle and homologues for the thioredoxin pathways (glutaredoxin, thioredoxin, and thioredoxin reductase), which are involved in various essential redox reaction in the cell) are also absent.9,10,19 A recent comparative genomic analysis of 16 different T. whipplei strains, including the Twist isolate, revealed that the coding sequences in this bacterium are highly conserved (ranging between 97.76% and 99.88%).20

17.1.2  Epidemiology Whipple’s disease has been reported to have affected 619  patients  (in addition to 79 unpublished cases) between the period 1907 to 1986.4 In 2008, the number of confirmed cases rose to 664.2 Whipple’s is a predominately a male disease (accounting for up to 86% of the reported cases).2,4,17 Age is another major factor in understanding the disease, and most cases only appear in patients between the ages of 40 and 60.2,4,17 Over the last few decades, the mean age of diagnosis of Whipple’s disease has been increasing, from 48.7 years (1965–1975), 50.7 years (1976–1985) to 57 years (1986–1995).2 Whipple’s disease is also likely to be race specific, as a majority of the patients are of European descent, chiefly in North America and Europe.2,4,17 Whipple’s disease is usually detected more frequently in farmers and carpenters, and the frequency of the disease is estimated to be less than one case per million of population per year.2,4 There has been no confirmed human-to-human transmission of disease.2 Furthermore, a limited number of PCR studies on a range of animals’ intestines suggested that none of the animals has T. whipplei present.17

17.1.3  Clinical Features Whipple’s disease is known as a multisystem disorder, with a majority of organs and organ systems being affected, although its effect on the gastrointestinal (GI) system is particularly notable.4 The manifestation of the “classic” Whipple’s disease includes arthralgia, weight loss, diarrhea, abdominal pain, and low-grade fever or chills. It may be years before all these clinical features of Whipple’s disease fully manifest in a patient.

Molecular Detection of Human Bacterial Pathogens

Symptoms that manifest involve the GI tract, the central nervous system (CNS), the joints, and the cardiovascular systems. Gastrointestinal Manifestations. GI symptoms of Whipple’s disease are usually described as weight loss, malabsorption, diarrhea, and abdominal pain, which may in turn progress to abdominal lymphadenopathy and a severe wasting syndrome. Weight loss is the most common manifestation (80%–90%) of Whipple’s disease, with most patients losing a net weight of between 5 and 15 kg per year. The most severe cases may lead to cachexia. Gastrointestinal bleeding is also common and may be caused by hematochezia. Furthermore, in some patients, the presenting intestinal lesion may appear similar to the lesions of untreated celiac disease. Stools samples from patients tend to be voluminous and fatty due to malabsorption.4,17,21 Central Nervous System Manifestations. Neurological manifestation is observed in 10%–40% of patients reported to have Whipple’s disease. Most often, neurological manifestations occur as a relapse after antibiotic treatment of intestinal and other manifestations of Whipple’s disease. CNS manifestation appearing with the intestinal disease is less frequent. However, approximately 50%–70% of patients are reported to have dementia, altered levels of consciousness, memory impairment, and confusion. Whipple’s disease affects ocular muscle movement; it is particularly present as progressive supranuclear ophthalmophegia with oculomasticatory myorhythemia or oculofacioskeletal myorhythemia. Although less well known, Whipple’s disease has been shown to produce slow rhythmical movement of facial and other muscles. Presence of abnormalities is obvious on neuroimaging scans, although there is no typical pattern or distribution to the CNS manifestations. Other neurological symptoms that may appear are epilepsy, focal cerebral lesion, ataxia seizures, headache, and meningitic features.4,17,22–26 Joint Manifestation. Articular symptoms account for the largest group (79%–90%), present in patients outside the GI manifestations. Whipple’s disease is harder to diagnosis in earlier stages because joint manifestations may precede GI or other systemic symptoms for several years. Arthragias and arthritis are the most common joint manifestations in patients, most of them occurring in knees, elbows, fingers, ankles, and shoulder joints.4,17,27 Cardiovascular System. Cardiovascular manifestations, including endo-, myo-, and pericarditis, appear in one-third of patients with Whipple’s disease. It was thought that endocarditis with major valve destruction was common (58%); however, with the use of PCR, a recent case study found that in 19 cases, T. whipplei infection was a factor in culture-negative endocarditis in Whipple’s disease.28,29 Other Manifestations. Another symptom of Whipple’s disease is the appearance of grayish-brown skin hyperpigmentation in about half of all cases. Ocular manifestations including uveitis, vitritis, retinitis, retrobulbar neuritis, and papilledema may also present; however, it is rare (5%). Pulmonary manifestations occur in 30%–40% of patients with Whipple’s disease, including pluritic chest pain, chronic cough, and dyspnea.4,17

Tropheryma

17.1.4  Diagnosis 17.1.4.1  Phenotypic Techniques Histopathological Examination. When Whipple’s disease—even if the primary presentation does not involve the GI tract—is suspected, the first approach of investigation is to undertake an upper GI endoscopy in which multiple biopsies from the lower duodenum and upper ileum are taken and examined by microscopy. Involvement of the intestine is seen in the majority of Whipple’s disease, so illnesses presenting with symptoms are rarely negative. However, approximately 10%–15% of cases may not show gastrointestinal symptoms and biopsies are negative, although PCR investigations of other organs may be positive. As such, biopsies of other organs may be useful.2 Because neurological signs occur in between 10% and 40% of patients with Whipple’s disease, periodic acid-Schiff (PAS) staining of cerebrospinal fluid or brain biopsies for PAS-positive inclusion may be informative in patients with CNS symptoms. Other biopsy samples that may be investigated include synovial fluid and/or tissues; soft tissue of patients with arthralgia or spondyodiscitis; cardiac valves of patients with culture negative endocarditis; bone marrow; or biopsies of the skin, liver, muscle, and eye.2 Microscopy and Electron Microscopy. Whipple’s disease was first described by George Whipple in 1907 and was first stained as sickled-form particules in macrophages using the periodic acid-Schiff stain in 1949. Periodic acid selectively oxidizes the glucose residues and creates aldehyde that reacts with the Schiff reagent, resulting in a purple magenta color reaction. The stain is used in histology to stain carbohydrate-rich macromolecules, such as glycogen, glycoprotein, and proteoglycans, commonly found in connective tissues and basal laminae. Detection of the bacterium with a stain predominantly used in histology and pathology was consistent with the observation of polysaccharide rich inner layer of its trilaminar cell wall by electron microscopy. The samples are typically digested with diastase to eliminate staining of glycogen in the biopsy samples, which may otherwise lead to a false-positive result.17 Electron microscopy examination indicated that the trilaminar layers consist of the aforementioned polysaccharide inner layer, an electron-translucent middle layer, and an electron-dense outer membrane that is somewhat similar to the membrane of a gram-negative bacterium that is absent of polysaccharides.17 Histopathologically, the bacterium can be detected in the foamy macrophages in the lamina propria of duodenal endoscopy biopsy or in the abdominal lymph nodes. PAS staining of the inclusion is not diagnostic of T.whipplei infection because other organisms, including yeast cells and fungal hyphae in tissues, may also be stained (http://www. hardydiagnostics.com/catalog2/hugo/Stains.htm). As such, other confirmatory tests are required. Culture of the Bacterium. T. whipplei was successfully cultivated and maintained in human embryonic fibroblast cell line (HEL) following shell vial centrifugation technique,

183

which was used with the intent of enhancing adhesion of the bacteria to the HEL cells7 after many previous failed attempts.15 The first strain, Twist-MarseilleT (=CNCM I-2202T), from an aortic valve of 42-year-old male diagnosed with endocarditis, was confirmed by PAS staining, amplification, and sequencing of the 16S rRNA gene and electron microscopy.6,7,30,31 Another strain (TW08/27) was isolated by a second group from cerebrospinal fluid of a women after presentation with severe weight loss, who grew the pathogen in murine fibroblast cell line, MRC-5, and primary human foreskin fibroblast cells.9,32,33 Up until 2003, the bacterium had only been successfully cultured in eukaryotic cells systems but the introduction of a cell-free axenic medium supplemented with amino acids enabled the faster isolation of T. whipplei.11 Specifically, the supernatant of T. whipplei–infected fibroblasts was mixed with DMEM/F12 medium and supplemented with fetal calf serum, l-glutamine, and human nonessential amino acids.11 Using the initial isolation methods of co-culturing the pathogen with eukaryotic cell lines, the doubling time of the bacterium ranged from between 4 and 17 days.7,9 In the cell-free axenic medium, the bacterium doubling time decreased to approximately 28 h.11 However, due to the difficulties associated with isolation and propagation of the bacterium, only two research laboratories in the world are involved in culturing the bacterium to date. Serological Assays. Antibodies to T. whipplei may be detected by western blot. The assay tests total immunoglobulins (IgT), including IgG, IgM, and IgA heavy and light chains. The production of rabbit polyclonal antibodies has also facilitated its detection in tissues by immunohistochemical staining.30,34–37 17.1.4.2  Genotypic Techniques A number of T. whipplei gene regions have been targeted for investigation in Whipple’s disease. These include the 16S rRNA operons, 16S-23S rRNA spacer region (or internal transcribed spacer), 5S rRNA, heat-shock protein 65 gene (hsp65 or groEL2).8,12,16,33,36,38–64 The 16S rRNA gene (16S rDNA) serves as a good tool for evolutionary analysis of bacteria strains because the gene accumulates mutations over time so that the phylogenetic distance of one bacterium from another can be calculated based on the mutational rates. However, some regions of the 16S rRNA gene are conserved among all bacteria. It was using these conserved regions that enabled confirmation of the etiologic agent for Whipple’s disease directly from DNA extracted from tissue.5,65 In a preliminary study, the 16S-23S rRNA intergenic spacer was reported to be homogenic,49 suggesting that it may be useful for diagnosing T. whipplei. However, a revised study of the same intergenic spacer region showed that this region is more hypermutable than its flanking structural genes (16S rRNA and 23S rRNA) and therefore is useful for species identification and genotyping purpose.49 The 16S-23S rRNA intergenic spacer serves as a useful target for genotyping T. whipplei strains, and seven different

184

spacer types have been identified so far; most spacers differ by only a few nucleotides along the 293–294 bp sequence.18 T. whipplei possesses several stretches of variable-number tandem repeat (VNTR) sequences; which can be also exploited for typing studies. After completion of the entire Tropheryma genome sequencing, other genes have been identified for molecular characterization of the bacterium. These include genes that encode the DNA-dependent RNA polymerase (rpoB), elongation factor TU (tuf), ATPase beta subunit (atpD), RNase P RNA (rnpb) and WiSP protein, and heat-shock protein 6. The rpoB gene, which is highly conserved in many bacteria, encodes the β subunit of the bacterial RNA polymerase, which catalyses the synthesis of RNA. The tuf gene, which contains a conserved P-loop motif of IGHVDHGKTT, encodes the elongation factor Tu that delivers aminoacyl-tRNA to A-site of the ribosomes during protein biosynthesis. The atpD gene is involved in F1F0-type ATP synthesis and is involved in regulating intracellular pH or synthesizing ATP in many enteric bacteria. The WiSP family of proteins is involved in surface protein interactions. The heat-shock protein 6 is implicated in the folding of proteins in prokaryote and eukaryotic cells and is required for normal cell growth. Being stress-induced, it acts to stabilize or protect disassembled polypeptides under heat-shock conditions. Using some of these genes as targets, Maiwald et al.8 identified the closest relatives phylogenetically for T.whipplei, which differ depending on the genes analyzed.18 For instance, the closest relatives for T. whipplei are Nocardioides jenseni (for rpnB), Propionibacterium acnes (for groEL2), Micrococcus luteus (for tuf), and S. coelicolor (for atpD and rpoB).18 Real-time quantitative PCR (qPCR) detection of T. whipplei is based on the amplification of the target sequence using specific primers and the recognition and hybridization of a probe specific for a target sequence in the DNA target, followed by the displacement of the probe as the target sequence is amplified.66 Fenollar et al.67 used quantitative PCR to investigate the frequency of T. whipplei infection in the feces of 150 healthy Senegalese children.

17.2  METHODS 17.2.1  Sample Preparation DNA from bronchoalveolar lavage (BAL) fluid sample may be extracted in a MagnaPure LC workstation (Roche Diagnostics) by using the MagnaPure LC DNA Isolation Kit II (Roche Diagnostics). Briefly, BAL sample (500 μL) is centrifuged in a 1.5 mL tube for 15 min at 14,000 × g, and 400 μL of the supernatant is discarded. The pellet is resuspended in the remaining 100 μL, mixed with 100 μL of lysis buffer and 50 μL of proteinase K, incubated overnight at 56°C, disrupted for 1 min with glass beads in a MagnaLyser (Roche Diagnostics), and then processed on the MagnaPure LC workstation according to the manufacturer’s recommendations.

Molecular Detection of Human Bacterial Pathogens

Alternatively, saliva (300 μL) is first liquefied with 300 μL of 3% NALC (N-acetyl-l-cysteine; Sigma), centrifuged for 10 min at 14,000 × g, and the supernatant discarded. Cerebrospinal fluid specimens, gastric aspirates, and rinse fluid of knee are centrifuged for 10 min at 14,000 × g, and supernatants discarded. The pellets are suspended in 200 μL of digestion buffer containing 200 μg of proteinase K per ml and incubated at 55°C for 1.5 h with agitation. DNA is extracted with QIAamp DNA binding columns (Qiagen) according to the manufacturer’s protocol except that in the final step only 100 μL (instead of 200 μL) of elution buffer AE is used. 5 μL of the eluate is used for PCR.68 Biopsy specimens are suspended directly in 200 μL of digestion buffer containing 200 μg of proteinase K per ml and incubated at 55°C for 1.5 h with agitation. DNA is extracted with QIAamp DNA binding columns as above. The paraffin-embedded tissue (two to five 5- to 10-μm-thick sections per specimen) is suspended in 200 μL of digestion buffer (50 mM Tris-HCl pH 8.5, 1 mM EDTA, 0.5% sodium dodecyl sulfate) without proteinase K. Mixtures are incubated at 95°C for 10 min with moderate agitation in a thermomixer (Eppendorf) followed by centrifugation at 14,000 × g for 30 min at room temperature. The resulting paraffin layers are removed with sterile pipette tips. Proteinase K (Roche Diagnostics) is then added to a final concentration of 200 μg per mL for further processing as for biopsy specimens.68 One loop of T. whipplei bacterial biomass is suspended in 200 μL of 4% CHELEX 100 (Bio-Rad). After incubation at 95°C for 15 min and centrifugation at 14,000 × g for 10 min, 1 μL of the supernatant is used for PCR.68

17.2.2  Detection Procedures (i) Nested PCR  Morgenegg et al.68 designed a nested PCR assay for identification of Tropheryma whipplei with primers from the heat-shock protein 65 gene (hsp65). In the first PCR, primers whipp-frw1 and whipp-rev generate a fragment of 500 bp; and in the second PCR, primers whipp-frw2 and whipp-rev generate a fragment of 357 bp from T. whipplei isolate (Table 17.1). Procedure 1. The first-round PCR mixture (50 μL) comprises 200 μM each of dATP, dCTP, dGTP, and dTTP, 25  pmol each of primers (whipp-frw1 and whipp-rev), 5 μL of 10× PCR buffer, 2.5 U of AmpliTaqGold polymerase (Perkin-Elmer), and 1 μL DNA template. 2. The first-round PCR mixture is subjected to an initial activation-denaturation step of 12 min at 95°C; 40 cycles of 1 min at 95°C, 57°C, and 72°C, respectively; and a final extension step of 10 min at 72°C, on a Gene Amp PCR System 9600 (Perkin-Elmer). 3. The resulting PCR product (10 μL) is separated by 1.5% agarose gel electrophoresis, stained with ethidium bromide, and visualized under UV light (a specific fragment of 500 bp is expected).

185

Tropheryma

TABLE 17.1 Primers for Specific Amplification of T. whipplei hsp65 Gene Primer whipp-frw1 whipp-rev whipp-frw2 whipp-rev a





Sequence (5′→3′)

Positiona

First-Round PCR TGACGGGACCACAACATCTG ACATCTTCAGCAATGATAAGAAGTT Second-Round PCR CGCGAAAGAGGTTGAGACTG As above

Amplicon Size (bp)

728–747 1231–1207

500

872–881 1231–1207

357

Corresponding to the hsp65 gene sequence of E. coli (GenBank accession no. X07850).

4. The second-round PCR mixture (50 μL) is made up of 200 μM each of dATP, dCTP, dGTP, and dTTP, 25 pmol each of primers (whipp-frw2 and whipp-rev), 5 μL of 10× PCR buffer, 2.5 U of AmpliTaqGold polymerase (Perkin-Elmer) and 1 μL of the firstround PCR product as template. Amplification is carried out with an initial 95°C for 12 min; 30 cycles of 1 min at 95°C, 60°C, and 72°C, respectively; and a final 72°C for 10 min. The amplified product is examined as above (a specific fragment of 357 bp is expected). 5. Optionally, the second-round PCR product (357 bp) may be verified by restriction enzyme digestion (RFLP analysis). Briefly, 10 μL PCR product is added with 2 μL of 10× restriction buffer, 1 μL of endonuclease XhoI (10 U in buffer H; Roche Diagnostics), and 7 μL of distilled water. The mixture is incubated at 37°C for 3 h. The digest (20 μL) and 10 μL of the undigested products are electrophoresed on a 3% agarose gel (2% NuSieveGTP agarose, FMC BioProducts, and 1% UltraPure agarose, Life Technologies), stained with ethidium bromide, and photographed under UV light. Three fragments of 202, 92, and 63 bp are expected from the digested product compared to the single fragment of 357 bp from the undigested product.

Note: The T. whipplei–specific seminested PCR assay, based on the partial hsp65 gene sequence, demonstrates high sensitivity and specificity. Seventeen patients shown to be positive by 16S rDNA PCR and/or internal transcribed spacer (ITS) PCR were also positive by the nested hsp65 PCR. To prevent possible crossover contamination from previous amplicons, dUTP instead of dTTP may be used in the first-round PCR along with 2% Tween 20 and uracil N-glycosylase (0.25U per 50 μL; Inotech, Dottikon, Switzerland). (ii) Quantitative PCR  Fenollar et al.69 outlined a testing scheme for identification of Tropheryma whipplei from clinical samples (e.g., 1 g stool, 200 μL saliva, 1 biopsy, or

200 μL body fluid) processed by QIAamp DNA MiniKit (Qiagen). The scheme involved use of two quantitative PCR (qPCR) with primers from the repetitive sequences of the bacterium. From October 2003 to March 2004, the T. whipplei–specific qPCR assay (targeting a 164 bp sequence) incorpo­rated primers 53.3F (5′-AGAGAGATGGGGTGCAGGAC-3′) and 53.3R (5′-AGCCTTTGCCAGACAGACAC-3′) in a FastStart DNA Master SYBR Green kit (Roche). If the result of this first assay was positive, it was systematically confirmed by a second qPCR using primers targeting a different gene region: 342F (5′-AGATGATGGATCTGCTTTCTTATCTG-3′) and 492R (5′-AACCCTGTCCTGCACCCC-3′). Since April 2004, the T. whipplei–specific qPCR (targeting a 155 bp sequence) incorporated the primers TW27F (5′-TGTTTTGTACTGCTTGTAACAGGATCT-3′) and TW182R (5′-TCCTGCTCTATCCCTCCTATC ATC-3′), and a TaqMan probe (27F–182R, 6-FAMAGAGATACATTTGTGTTAGTTGTTACA-TAMRA). If the result of this first assay was positive, it was systematically confirmed by a second PCR using primers that targets a different 150  bp region: TW13F (5′-TGAGTGATGGTAGTCTGAGAGATATGT-3′) and TW163R (5′-TCCATAACAAAGACAACAACCAATC-3′) together with a TaqMan probe (13F–163R, 6-FAMAGAAGAAGATGTTACGGGTTG-TAMRA). (iii) Quantitative Real-Time PCR  Bousbia et al.70 described a quantitative real-time PCR using the QuantiTect Probe PCR Kit on LightCycler instrument (Roche Diagnostics) with cycling conditions specified by the manufacturer. The specimens are first tested with the primers Twhi3F: 5′-TTGTGTATTTGGTATTAGATGAAACAG-3′ and Twhi3R: 5′-CCCTACAATATGAAACAGCCTTTG-3′ and the specific TaqMan probe Twhi3: 6-FAMGGGATAGAGCAGGAGGTGTCTGTCTGG-TAMRA. When the specimen is positive in the above assay, the result is confirmed by a second quantitative PCR with the primers Twhi2F: 5′-TGAGGATGTATCTGTGTATGGGACA-3′ and Twhi2R: 5′-TCCTGTTACAAGCAGTACAAAA CAAA-3′ and the Twhi2 probe: 6-FAM-GAGAGATGGGG TGCAGGACAGGG-TAMRA.

186

Molecular Detection of Human Bacterial Pathogens

(iv) 16S rRNA Sequencing  Bousbia et al.70 utilized a broad-range primer-based PCR targeting the 16S rRNA gene followed by sequencing to identify Tropheryma whippeli associated with pneumonia. The eubacterial broad-range 16S rRNA primers for amplification consist of 536F: 5′-CAGCAGCCGCGGTAATAC-3′ and rp2: 5′-ACGGCTACCTTGTTACGACTT-3′. Procedure 1. The PCR mixture (50-μL) is composed of 5 μL extracted DNA, 1 × PCR buffer (5 μL), 2 μL of 25 mM MgCl2, 200 μM of each dNTP, 0.2 mM of each 536F and rp2 primer, and 1 U of HotStar Taq DNA Polymerase (Qiagen). 2. Amplification is performed with an initial denaturation and activation step at 95°C for 15 min; 35 cycles of 95°C for 1 min, 62°C for 30 s, and 72°C for 90 s; and an extension step at 72°C for 10 min, in an ABI themocycler (Applied Biosystems). 3. The PCR products are purified by using the NucleoFast 96 PCR Kit (Macherey-Nagel), and 4 μL of purified PCR product is sequenced in 20 μL final volume containing sequencing buffer, 3.2 pmol of forward (536F) or reverse (rp2) primer, 3 μL of Big Dye Terminator V1.1 mix (Applied Biosystems), and 8 μL of deionized water with cycling conditions specified by the manufacturer. 4. Sequencing reactions are then purified using Sephadex Gel-Filtration (Sigma-Aldrich), and the purified products are sequenced on an ABI PRISM 3130xl genetic analyzer (Applied Biosystems). Sequences obtained are analyzed with Autoassembler software (Perkin-Elmer) and compared with those available in the GenBank database using the BLAST program (www.ncbi.nlm.nih.gov/ BLAST). Note: When the BAL fluid is polymicrobial, 3 μL of purified PCR product (from step 3) is cloned into PCR II TA cloning

vector (Invitrogen) according to the manufacturer, and 56–64 white colonies are screened for each specimen. The universal M13 primers (M13F: 5′-GTAAAACGACGGCCAG-3′ and M13R: 5′-CAGGAAACAGCTATGAC-3′) are then employed to sequence the cloned inserts from both directions. (v) Genotyping  Li et al.71 developed a genotyping system for Tropheryma whipplei on the basis of four highly variable genomic sequences (HVGSs). The HVGS 1 marker separates T. whipplei into eight genotypes (1–8); the HVGS 2 marker also separates T. whipplei into eight genotypes (1–8); the HVGS 3 marker into six genotypes (1–6); and the HVGS 4 marker into four genotypes (1–4) (Table 17.2). Procedure 1. PCR mixture (25 µL) is made up of 5 µL of 3 ng/µL DNA, 50 pM each primer, 200 µM (each) dNTPs, 1.5 U Hotstart Taq DNA polymerase (Qiagen), 2.5 µL 10× PCR buffer; and 1 µL 25 mM MgCl2. Sterile water is used as a negative control in each PCR run. 2. The mixture is subjected to an initial 95°C for 15 min; 40 cycles of 94°C for 30 s, 55°C for 30 s, and 72°C for 1 min; and a final 72°C for 5 min, in a PTC-200 automated thermal cycler (MJ Research). 3. PCR products are purified by using the MultiScreen PCR filter plate (Millipore). Amplicons are sequenced in both directions using BigDye 1.1 chemistry (Applied Biosystems) on an ABI 3130xl automated sequencer (Applied Biosystems). All sequences are checked twice in both directions to ensure the reliability of the typing method. 4. Nucleotide sequences are edited using the Autoassembler package (Perkin-Elmer). Multiple alignments of sequences are carried out using the CLUSTALW software. For phylogenetic analysis, sequences of the four HVGSs are concatenated. Phylogenetic relationships among T. whipplei genotypes are inferred using the unweighted pair group

TABLE 17.2 Primers for Amplification and Sequencing of Tropheryma whippeli Highly Variable Genomic Sequences (HVGSs) HVGS HVGS 1 HVGS 2 HVGS 3 HVGS 4

Content of HVGS

Genotype

Size (bp)

TWT133 and intergenic spacer (TWT133-TWT134) proS/prolyl-tRNA synthetase (TW183) secA-hp (TWT131) intergenic spacer TW183 and intergenic spacer (TWT183-TWT184)

8

227

8

318

6

150

4

162

Source:  Adapted from Li, W. et al., Microbiology, 154, 521, 2008.

Forward and Reverse Primers (5′–3′) GCTGCGCGAAGTAATTTG AGATACATGCGGAGATACT GCCTTGACTATGACATAATCAA TCGGACTAAAAGTGCGACAC TTTGTCATAGGCATTTCTGTAG AGACCTCACTGTTATACGGAT CGGATCTTCACGAAATGTCC ATAACAAGAAGCTGGATATGC

Tropheryma

method with arithmetic mean (UPGMA), neighborjoining and maximum-parsimony methods within the MEGA 3.1 software. Note: For each HVGS, a sequence type is defined as a sequence exhibiting unique mutation(s). HVGS genotypes are defined as unique combinations of the four HVGS types.

17.3  CONCLUSION Tropheryma whipplei is a gram-positive, nonmotile bacterium that is responsible for Whipple’s disease—a systemic chronic infection displaying several clinical traits, including histological lesions in the gastrointestinal tract, endocarditis with negative blood cultures, and isolated neurological infection.72,73 The disease predominantly affects middleaged men. Although Whipple’s disease is rare, it can be fatal without antibiotic treatment. Because the culture is notably time consuming and has only been successfully achieved in two laboratories, the current diagnostic scheme for Whipple’s disease relies on diastase-resistant periodic acid-Schiff staining together with PCR, involving organ/tissue biopsy (small intestine, cardiac valve, and cerebrospinal fluid) as well as saliva and stool specimens.74 When classic Whipple’s disease with gastrointestinal symptoms is suspected, qPCR screening of saliva and stool specimens provides a valuable noninvasive way to confirm the infection, especially if bacterial load in stool is >104 CFU/g. For localized Whipple’s disease (often accounting for nearly one-quarter of cases), use of broad-spectrum and specific qPCR assay is essential for diagnosis.69

REFERENCES



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187 11. Renesto, P. et al., Genome-based design of a cell-free culture medium for Tropheryma whipplei, Lancet, 362, 447, 2003. 12. Drancourt, M., Carlioz, A., and Raoult, D., rpoB sequence analysis of cultured Tropheryma whippelii, J. Clin. Microbiol., 39, 2425, 2001. 13. Marth, T., and Raoult, D., Whipple’s disease, Lancet, 361, 239, 2003. 14. Silva, M.T., Macedo, P.M., and Moura Nunes, J.F., Ultrastructure of bacilli and the bacillary origin of the macrophagic inclusions in Whipple’s disease, J. Gen. Microbiol., 131, 1001, 1985. 15. Dobbins, W.O. 3rd, Whipple’s disease: An historical perspective, Q. J. M., 56, 523, 1985. 16. Maiwald, M. et al., Tropheryma whippelii DNA is rare in the intestinal mucosa of patients without other evidence of Whipple disease, Ann. Intern. Med., 134, 115, 2001. 17. Dutly, F., and Altwegg, M., Whipple’s disease and “Tropheryma whippelii,” Clin. Microbiol. Rev., 14, 561, 2001. 18. Maiwald, M. et al., Organization, structure, and variability of the rRNA operon of the Whipple’s disease bacterium (Tropheryma whippelii), J. Bacteriol., 182, 3292, 2000. 19. Crapoulet, N. et al., Tropheryma whipplei genome at the beginning of the post-genomic era, Curr. Genom., 6, 195, 2005. 20. La, M.V. et al., Comparative genomic analysis of Tropheryma whipplei strains reveals that diversity among clinical isolates is mainly related to the WiSP proteins, BMC Genomics, 8, 349, 2007. 21. Amsler, L. et al., Prevalence of Tropheryma whipplei DNA in patients with various gastrointestinal diseases and in healthy controls, Infection, 31, 81, 2003. 22. Rickman, L.S. et al., Uveitis caused by Tropheryma whippelii (Whipple’s bacillus), N. Engl. J. Med., 332, 363, 1995. 23. Louis, E.D. et al., Diagnostic guidelines in central nervous system Whipple’s disease, Ann. Neurol., 40, 561, 1996. 24. Helliwell, T.R. et al., Dermatomyositis and Whipple’s disease, Neuromuscul. Disord., 10, 46, 2000. 25. Messori, A. et al., An unusual spinal presentation of Whipple disease, Am. J. Neuroradiol., 22, 1004, 2001. 26. Matthews, B.R. et al., Cerebellar ataxia and central nervous system Whipple disease, Arch. Neurol., 62, 618, 2005. 27. Ayoub, W.T. et al., Bone destruction and ankylosis in Whipple’s disease, J. Rheumatol., 9, 930, 1982. 28. Richardson, D.C. et al., Tropheryma whippelii as a cause of afebrile culture-negative endocarditis: The evolving spectrum of Whipple’s disease, J. Infect., 47, 170, 2003. 29. Marin, M., Tropheryma whipplei infective endocarditis as the only manifestation of Whipple’s disease, J. Clin. Microbiol., 45, 2078, 2007. 30. Raoult, D. et al., Culture and immunological detection of Tropheryma whippelii from the duodenum of a patient with Whipple disease, JAMA, 285, 1039, 2001. 31. Raoult, D., Fenollar, F., and Birg, M.L., Culture of T whipplei from the stool of a patient with Whipple’s disease, N. Engl. J. Med., 355, 1503, 2006. 32. Drancourt, M. et al., Culture of Tropheryma whippelii from the vitreous fluid of a patient presenting with unilateral uveitis, Ann. Intern. Med., 139, 1046, 2003. 33. Fenollar, F., and Raoult, D., Molecular techniques in Whipple’s disease, Expert Rev. Mol. Diagn., 1, 299, 2001. 34. Baisden, L. et al., Diagnosis of Whipple disease by immunohistochemical analysis: A sensitive and specific method for the detection of Tropheryma whipplei (the Whipple bacillus) in paraffin-embedded tissue, Am. J. Clin. Pathol., 118, 742, 2002.

188 35. Lepidi, H. et al., Whipple’s disease: Immunospecific and quantitative immunohistochemical study of intestinal biopsy specimens, Hum. Pathol., 34, 589, 2003. 36. Lepidi, H. et al., Cardiac valves in patients with Whipple endocarditis: Microbiological, molecular, quantitative histologic, and immunohistochemical studies of 5 patients, J. Infect. Dis., 190, 935, 2004. 37. Kowalczewska, M. et al., Identification of candidate antigen in Whipple’s disease using a serological proteomic approach, Proteomics, 6, 3294, 2006. 38. Lowsky, R. et al., Diagnosis of Whipple’s disease by molecular analysis of peripheral blood, N. Engl. J. Med., 331, 1343, 1994. 39. Maiwald, M. et al., Detection of Tropheryma whippelii DNA in a patient with AIDS, J. Clin. Microbiol., 33, 1354, 1995. 40. Cohen, L. et al., Polymerase chain reaction of cerebrospinal fluid to diagnose Whipple’s disease, Lancet, 347, 329, 1996. 41. Maiwald, M. et al., Reassessment of the phylogenetic position of the bacterium associated with Whipple’s disease and determination of the 16S-23S ribosomal intergenic spacer sequence, Int. J. Syst. Bacteriol., 46, 1078, 1996. 42. von Herbay, A., Ditton, H.J., and Maiwald, M. Diagnostic application of a polymerase chain reaction assay for the Whipple’s disease bacterium to intestinal biopsies, Gastroenterology, 110, 1735, 1996. 43. Dauga, C., Miras, I., and Grimont, P.A. Strategy for detection and identification of bacteria based on 16S rRNA genes in suspected cases of Whipple’s disease, J. Med. Microbiol., 46, 340, 1997. 44. Ramzan, N.N. et al., Diagnosis and monitoring of Whipple disease by polymerase chain reaction, Ann. Intern. Med., 126, 520, 1997. 45. Celard, M. et al., Polymerase chain reaction analysis for diagnosis of Tropheryma whippelii infective endocarditis in two patients with no previous evidence of Whipple’s disease, Clin. Infect. Dis., 29, 1348, 1999. 46. Delanty, N. et al., Synovial fluid polymerase chain reaction as an aid to the diagnosis of central nervous system Whipple’s disease, Ann. Neurol., 45, 137, 1999. 47. Ehrbar, H.U. et al., PCR-positive tests for Tropheryma whippelii in patients without Whipple’s disease, Lancet, 353, 2214, 1999. 48. Hinrikson, H.P. et al., Detection of three different types of “Tropheryma whippelii” directly from clinical specimens by sequencing, single-strand conformation polymorphism (SSCP) analysis and type-specific PCR of their 16S-23S ribosomal intergenic spacer region, Int. J. Syst. Bacteriol., 49, 1701, 1999. 49. Hinrikson, H.P., Dutly, F., and Altwegg, M. Homogeneity of 16S-23S ribosomal intergenic spacer regions of Tropheryma whipplei in Swiss patients with Whipple’s disease, J. Clin. Microbiol., 37, 152, 1999. 50. Hinrikson, H.P., Dutly, F., and Altwegg, M., Analysis of the actinobacterial insertion in domain III of the 23S rRNA gene of uncultured variants of the bacterium associated with Whipple’s disease using broad-range and “Tropheryma whippelii”-specific PCR, Int. J. Syst. Evol. Microbiol., 50, 1007, 2000. 51. Hinrikson, H.P., Dutly, F., and Altwegg, M. Evaluation of a specific nested PCR targeting domain III of the 23S rRNA gene of “Tropheryma whippelii” and proposal of a classification system for its molecular variants, J. Clin. Microbiol., 38, 595, 2000. 52. Lange, U., and Teichmann, J., Diagnosis of Whipple’s disease by molecular analysis of synovial fluid, Arthritis Rheum., 42, 1777, 1999.

Molecular Detection of Human Bacterial Pathogens 53. Misbah, S.A. et al., Anonymous survey of blood donors by polymerase chain reaction for Tropheryma whippelii, Q. J. M., 92, 61, 1999. 54. Pron, B. et al., Diagnosis and follow-up of Whipple’s disease by amplification of the 16S rRNA gene of Tropheryma whippelii, Eur. J. Clin. Microbiol. Infect. Dis., 18, 62, 1999. 55. Street, S., Donoghue, H.D., and Neild, G.H., Tropheryma whippelii DNA in saliva of healthy people, Lancet, 354, 1178, 1999. 56. Fenollar, F. et al., Quantitative detection of Tropheryma whipplei DNA by real-time PCR, J. Clin. Microbiol., 40, 1119, 2002. 57. Fenollar, F. et al., Culture of Tropheryma whipplei from human samples: A 3-year experience (1999 to 2002), J. Clin. Microbiol., 41, 3816, 2003. 58. Fenollar, F. et al., Use of genome selected repeated sequences increases the sensitivity of PCR detection of Tropheryma whipplei, J. Clin. Microbiol., 42, 401, 2004. 59. Fredricks, D.N., and Relman, D.A., Localization of Tropheryma whippelii rRNA in tissues from patients with Whipple’s disease, J. Infect. Dis., 183, 1229, 2001. 60. Geißdörfer, W. et al., Detection of a new 16S-23S rRNA spacer sequence variant (type 7) of Tropheryma whippelii in a patient with prosthetic aortic valve endocarditis, Eur. J. Clin. Microbiol. Infect. Dis., 20, 762, 2001. 61. Bruhlmann, P., Michel, B.A., and Altwegg, M., Diagnosis and therapy monitoring of Whipple’s arthritis by polymerase chain reaction, Rheumatology, 39, 1427, 2000. 62. Maibach, R.C., Dutly, F., and Altwegg, M., Detection of Tropheryma whipplei DNA in feces by PCR using a target capture method, J. Clin. Microbiol., 40, 2466, 2002. 63. Puéchal, X. et al., Polymerase chain reaction testing for Tropheryma whippelii in unexplained isolated cases of arthritis, Arthritis Rheum., 44, 1130, 2002. 64. Rolain, J.M., Fenollar, F., and Raoult, D., False positive PCR detection of Tropheryma whipplei in the saliva of healthy people, BMC Microbiol., 7, 48, 2007. 65. Wilson, K.H. et al., Phylogeny of Whipple’s disease associated bacterium, Lancet, 338, 474, 1991. 66. Sloan, L.M., Rosenblatt, J.E., and Cockerill, III, F.R., Detection of Tropheryma whipplei DNA in clinical specimens by LightCycler real-time PCR, J. Clin. Microbiol., 43, 3516, 2005. 67. Fenollar, F. et al., Tropheryma whipplei in Fecal Samples from Children, Senegal, Emerg. Infect. Dis., 15, 922, 2009. 68. Morgenegg, S., Dutly, F., and Altwegg, M., Cloning and sequencing of a part of the heat shock protein 65 gene (hsp65) of “Tropheryma whippelii” and its use for detection of “T. whippelii” in clinical specimens by PCR, J. Clin. Microbiol., 38, 2248, 2000. 69. Fenollar, F. et al., Value of Tropheryma whipplei quantitative polymerase chain reaction assay for the diagnosis of Whipple disease: usefulness of saliva and stool specimens for first-line screening, Clin. Infect. Dis., 47, 659, 2008. 70. Bousbia, S. et al., Tropheryma whipplei in patients with pneumonia, Emerg. Infect. Dis., 16, 258, 2010. 71. Li, W. et al., Genotyping reveals a wide heterogeneity of Tropheryma whipplei, Microbiology, 154, 521, 2008. 72. Fenollar, F., Puechal, X., and Raoult, D., Whipple’s disease, N. Engl. J. Med., 356, 55, 2007. 73. Fleming, J.L., Wiesner, R.H., and Shorter, R.G., Whipple’s disease: Clinical, biochemical, and histopathologic features and assessment of treatment in 29 patients, Mayo Clin. Proc., 63, 539, 1988. 74. Kowalczewska, M., and Raoult, D., Advances in Tropheryma whipplei research: The rush to find biomarkers for Whipple’s disease, Future Microbiol., 2, 631, 2007.

18 Tsukamurella Mireille M. Kattar CONTENTS 18.1 Introduction...................................................................................................................................................................... 189 18.1.1 Classification, Morphology, and Epidemiology.................................................................................................... 189 18.1.2 Clinical Features and Pathogenesis...................................................................................................................... 190 18.1.3 Diagnosis...............................................................................................................................................................191 18.1.3.1 Conventional Techniques........................................................................................................................191 18.1.3.2 Molecular Techniques............................................................................................................................ 192 18.2 Methods............................................................................................................................................................................ 194 18.2.1 Sample Preparation............................................................................................................................................... 194 18.2.2 Detection Procedures............................................................................................................................................ 196 18.3 Conclusions and Future Perspectives............................................................................................................................... 197 References.................................................................................................................................................................................. 197

18.1  INTRODUCTION The genus Tsukamurella was created in 1988 in honor of Michio Tsukamura (a Japanese microbiologist) to accommodate mesophilic gram-positive weakly acid fast aerobic actinomycetes with highly unsaturated mycolic acids in their cell walls, encompassing chains 62–78 carbon atoms in length. Their predominant menaquinone is of the MK-9 type. It currently includes 10 validly named species, among which five have been associated with human infections, whereas the other five have been solely isolated from environmental sources.

18.1.1  Classification, Morphology, and Epidemiology Classification and Epidemiology. The genus Tsukamurella belongs to the family Tsukamuralleceae in the suborder Corynebacterineae. The suborder Corynebacterineae belongs to the order Actinomycetales within the class Actinobacteria.1 Tsukamurellae are soil organisms and have been found in treated and untreated drinking water.2 The genus Tsukamurella was first established in 1988 to accommodate a group of chemically unique organisms characterized by a series of very long chain (62–78 carbon atoms), highly unsaturated (2–7 double bonds) mycolic acids, in addition to possessing meso-diaminopimelic acid, arabinose, and galactose, which are common to the genus Corynebacterium.3 The latter features are characteristic of the cell-wall chemotype IV.4 The type species Tsukamurella paurometabola, described by Steinhaus in 1941 as Corynebacterium

paurometabolum, was originally isolated from the mycetomes and ovaries of bed bugs (Cimex lectularius).5 In a comprehensive numerical phenetic study by Jones et al.,6 the type strain C. paurometabolum ATCC 8368 was found to be distinct from Corynebacterium species. The first reported human isolate of Tsukamurella, named Gordona aurantiaca, was identified in the sputum of a patient with tuberculosis, but its causal role in the patient’s disease was doubtful.7 In 1985, G. aurantiaca was renamed Rhodococcus aurantiacus by Tsukamura and assigned a new type strain, ATCC 25938.8 Chemical and numerical taxonomic studies of C. paurometabolum and G. aurantiaca revealed that they had similar cellular composition.9,10 Both possess highly unsaturated mycolic acids containing 62–78 carbon atoms. They also contain fully unsaturated menaquinones, which include MK-8, MK-9, and MK-10, with predominant 9 isoprene units (MK-9). This is in contrast to other mycolic acid-containing taxa such as Mycobacterium, Nocardia, Rhodococcus, and Corynebacterium species, which possess dihydrogenated menaquinones.11 Menaquinones are present in the cell membranes of most bacteria, where they function in electron transport, oxidative phosphorylation, and perhaps active transport. Based on these results and the more than 99% similarity in their 16S rRNA gene sequences, Collins et al. proposed in 1988 that C. paurometabolum and G.  aurantiaca be reduced to a single species and assigned to a new genus, Tsukamurella.3 The type species is Tsukamurella paurometabolum, and strain ATCC 8368 was designated as the type strain. Yassin later correctly renamed T. paurometabolum as T. paurometabola.12 Four other species associated with human infections were subsequently described. Two strains 189

190

of T. inchonensis were initially isolated from multiple blood cultures of a patient who had ingested hydrochloric acid and from the lung tissue of a patient who had a lung tumor, respectively.12 T. pulmonis was first isolated from the sputum of a 92-year-old woman diagnosed three years previously with pulmonary tuberculosis.13 Strains of T. tyrosinosolvens were initially recovered from elderly patients; two patients had cardiac pacemaker implants, and the organism grew from multiple blood cultures; in another two patients with chronic lung infections, the organisms were obtained from sputum cultures.14 T. strandjordii was isolated from multiple blood cultures of a 5-year-old child with acute myelogenous leukemia and an indwelling catheter.15 Five other species of Tsukamurella were proposed from 2003 to 2009, all of which were discovered in environmental samples. T. sunchonensis16 was isolated from activated sludge foam in Korea, and T. spumae17 and T. pseudospumae18 in the United Kingdom. In this respect, Tsukamurella constitute one of several genera of aerobic actinomycetes that have been associated with sludge foams. The activated sludge process is the most commonly used method for the treatment of wastewater. Activated sludge foams may represent a public health threat because of the potential spread of pathogens through aerosols.19 T. spongiae was isolated from a deep-water marine sponge off the coast of Curaçao in the Netherlands Antilles.20 The most recently described species is T. carboxydivorans, which was isolated from a soil sample collected from a roadside in Seoul, Korea.21 The Guanine + Cytosine content of Tsukamurella species ranges from 67% to 77% mol, with the highest G + C content observed in T. carboxydivorans. In contrast to the other species, T. carboxydivorans contains the longest chain mycolic acids with 81–95 carbon atoms, whereas T. spongiae contains the shortest with 58–75 carbon atoms. Morphology. Tsukamurella species are gram-positive, long, straight to slightly curved rod-shaped and partially acid alcohol fast measuring 0.5–0.8 µm by 1–5 µm.12–14 Acid fastness requires use of a modified acid-fast stain with a weaker decolorizer (1% sulfuric acid H2SO4). Clinical isolates have occasionally been reported to be acid-fast negative.22 On scanning electron microscopy, bacterial cells fragment into three to four parts, which separate and grow as independent cells. Growth is obligately aerobic. Visible colony growth from clinical specimens and dilute inocula occurs within 2 days. Growth in blood culture bottles from clinical specimens occurs within 1–2 days. Colonies show rudimentary to no branching. The cells are nonmotile and do not form spores, capsules, or aerial hyphae. All Tsukamurella species usually grow at temperatures ranging from 25°C to 35°C. T. paurometabola and T. pseudospumae also grow at 10°C, and T. inchonensis grows at 42°C. The optimal growth temperature for T. carboxydivorans is 30°C. After 24–48 h of incubation, colony size is 2–5 mm, and colonies are easily emulsified. Depending on the species, colony color is either cream white or pigmented. Colonies of T. paurometabola type strain are convex dry creamy white with entire edges. Colonies of T. pulmonis, T. strandjordii, T. spongiae, and T. carboxydivorans are also cream-colored rough with fringed

Molecular Detection of Human Bacterial Pathogens

edges and raised, wrinkled centers. Colonies of T. inchonensis and T. tyrosinosolvens are also rough and dry, with fringed edges, but are yellow to orange with no diffusible pigment. Colonies of T. sunchonensis, T. spumae, and T. pseudospumae have irregular margins and are orange to red.

18.1.2  Clinical Features and Pathogenesis The pathogenic role of Tsukamurellae was initially a subject of debate.23,24 Tsukamura and Mizuno first reported that since 1971, they had isolated Gordona aurantiaca from sputa of 14 patients with tuberculosis or “tuberculosis-like” illness.7,25 They stated, however, that this organism had not caused disease in humans. In 1982, Tsukamura and Kawakami reported the first case of a lung infection caused by Gordona aurantiaca that mimicked tuberculosis.25 Since then, 24 case reports and six short case series of human infection have been published, and these cases occurred nearly equally in male and female patients.22,26–51 Most cases occurred in adult patients, but there were three cases in pediatric patients.37,52 These infections occurred predominantly in immunocompromised patients and, occasionally, in immunocompetent hosts. Tsukamurella species appear to have a low pathogenic potential and have been mostly linked to opportunistic infections in patients with foreign medical devices, which account for over 80% of cases reported. The majority of cases consisted of bloodstream infections secondary to central venous catheters placed several months earlier for treatment of cancer.30,34,36,37,40,45,48 The organisms were frequently cultured from the catheter tips and peripheral blood, as well as from blood cultures drawn through the central catheters. Infections sometimes accompanied a febrile neutropenic episode. None of the patients had tunnel or catheter exit-site infection. In virtually all these cases, positive blood cultures persisted despite treatment with various antibiotics, thereby necessitating the removal of the catheter, which resulted in the patient’s prompt recovery and subsequently negative blood cultures. No firm conclusions can be drawn concerning the efficacy of antibiotics because therapy varied widely in terms of class and duration (reviewed in Ref. 48). Five cases occurred in patients with chronic renal failure.22,27,37,38,42 Two of these patients had indwelling catheters for hemodialysis. Another two patients had peritonitis secondary to peritoneal dialysis catheters. The organism was grown from blood or peritoneal fluid in all patients. It also grew from the catheter tip in one patient. In all five patients, initial antibiotic therapy with one or multiple drugs failed to control the infection, which only resolved upon removal of the catheter. Three patients had cardiac pacemaker implants.14,41 One patient had a chronic wound infection following prosthetic knee replacement, which resolved after 2 months of treatment with clarithromycin and ciprofloxacin, to which the organism was susceptible.35 The second most common syndrome associated with Tsukamurellae is pulmonary infection mimicking tuberculosis. Ten such cases have been reported, 70% of which were in patients with no evident predisposing factor.13,14,25,29,39,43,44,46,47

191

Tsukamurella

The organism was often grown repeatedly from sputum and, less commonly, from lung tissue. Two patients had lung cancer, and one patient was HIV positive.12,39,43 Treatment and follow-up information was available for four patients. Three patients responded to a combination of a quinolone and a rifamycin (rifampin or rifabutin) for 3–6 months, and one patient achieved cure following 4 months of treatment with trimethoprim and sulfamethoxazole. In one report, a HIVpositive female patient was colonized in the sputum with Tsukamurella species but did not develop any respiratory symptoms after a follow-up period of 4 months.53 Three cases of skin and soft tissue infection due to Tsukamurellae have been reported in immunocompetent adult patients. One case occurred in a 30-year-old female patient following surgical treatment for constrictive tenosynovitis.28 She developed a wound infection with multiple abscesses and arm gangrene that failed to respond to a combination of a quinolone, rifampin, and minocycline and eventually required amputation of her arm. Another patient had a chronic superficial infection that responded to topical hexamidine and fusidic acid.32 There have been two reported cases of central nervous system infection due to Tsukamurellae. One case of fatal meningitis occurred in a 43-year-old male patient with hairy cell leukemia.26 A 36-year-old female patient developed skull osteomyelitis and a brain abscess following purulent chronic otitis media.49 The infection responded to a 6-week combination therapy with amikacin and imipenem, followed by oral ciprofloxacin, clarithromycin, and thrimetroprimsulfamethoxazole. Lastly, Tsukamurellae species have been associated with ocular infections in immunocompetent adults in China. Three patients had mild conjunctivitis that responded to a 10-day course of topical chloramphenicol or polymyxin B and neomycin.51 Two patients developed keratitis, which responded to topical gentamicin or levofloxacin for 10 days to 3 weeks. Both patients had microtrauma to the corneal surface, which predisposed them to developing a deeper eye infection.50 In 1992, Auberbach et al. reported an outbreak of pseudoinfection with T. paurometabola involving 10 patients that occurred at a single hospital in South Carolina over a 30-month period.23 These isolates were grown from eight tissue specimens, a bronchial washing, and swab from a fistula. The isolates from the outbreak were similar by ribotyping. The specimens from the outbreak were more likely to have been processed in the mycobacteriology laboratory and to have been handled by one technician. Prior to 1995, infections with Tsukamurellae were all attributed to T. paurometabola, previously known as Gordona aurantiaca or Rhodococcus aurantiacus. However, the organism descriptions in most of these reports, including the pseudooutbreak, did not fit that of T. paurometabola type strain but rather strain ATCC 25938, which has been demonstrated to be a misnamed T. inchonensis strain.15 Following the original descriptions of T. inchonensis, T. pulmonis, and T. tyrosinosolvens from 1995 to 1997, and T. strandjordii in 2001, all reported human infections where

species identification were performed, were attributed to either T. pulmonis or T. tyrosinosolvens. Species identification of the isolates was achieved by phenotypic methods or 16S rRNA gene sequencing, or a combination of the two methods (see below). In terms of antimicrobial susceptibility patterns, and prior to the publication of the Clinical and Laboratory Standards Institute (CLSI) in vitro susceptibility testing guidelines for aerobic actinomycetes in 2003, testing was not standardized. Prior to 2003, published reports included such methods as disk diffusion, broth microdilution, broth macrodilution, E-test, and agar dilution. The reference method recommended by CLSI is broth microdilution.54 The primary antibiotic panel includes amikacin, tobramycin, amoxicillin-clavulanic acid, ceftriaxone, ciprofloxacin, clarithromycin, imipenem, linezolid, minocycline, and trimetroprim-sulfamethoxazole. The secondary antibiotic panel includes cefepime, cefotaxime, doxycycline, and gentamicin. Tsukamurellae are resistant in vitro to agents used for the treatment of tuberculosis, which include streptomycin, isoniazid, ethambutol, cycloserine, protionamide, and capreomycin.12–14 They are also resistant to a wide range of antibiotics of different classes. The mechanisms of antimicrobial resistance in Tsukamurellae are unknown. When grown on some hydrophobic carbon sources, they may produce glycolipids that are secreted but are also found attached to the bacterial cell wall. These glycolipids are believed to have antimicrobial properties.55 They are usually susceptible to quinolones, amikacin, clarithromycin, trimethoprimsulfamethoxazole, minocycline, and rifampin. They are reportedly resistant to penicillin, ampicillin, ampicillin/sulbactam, amoxicillin/clavulanic acid, carbenicillin, piperacillin/tazobactam, oxacillin, first-generation cephalosporins, cefoxitin, moxalactam, and vancomycin. They may be either susceptible or resistant to erythromycin, clindamycin, doxycycline, gentamicin, tobramycin, third-generation cephalosporins, imipenem, and chloramphenicol. Histotological examination of tissues infected with Tsukamurellae usually shows granulomatous inflammation without central necrosis, reminiscent of the tissue response encountered in some cases of tuberculosis.28,32 This has been attributed in a mouse model to the secretion of interferon-γ by recruited CD4+ T lymphocytes.56

18.1.3  Diagnosis 18.1.3.1  Conventional Techniques The morphological features on Gram staining, positive modified acid-fast staining, strictly aerobic growth, and colonial morphology represent the first indication that an organism belongs to the aerobic actinomycetes. Using phenotypic methods, members of the genus Tsukamurella may be confused with, and have been variously misidentified, as rapidly growing Mycobacterium species, Nocardia, Rhodococcus, or Corynebacterium species.15,22,30,32,36,57 Although Tsukamurella species are considered to be weakly positive on acid-fast staining, some original descriptions

192

state that some of the members may appear strongly acid fast.12 In addition, they grow on media used in the clinical laboratory for the isolation of Mycobacterium species, including Lowenstein-Jensen and Middlebrook 7H11 and similar to Mycobacterium species, are resistant to lysozyme. Furthermore, they frequently react in the same conventional phenotypic tests used for the identification of Mycobacterium species.37 Tsukamurella species also grow on sheep blood agar, chocolate, and MacConkey without crystal violet. Commercially available panels have limited value for identification of aerobic actinomycetes, which are generally not included in the database of these systems. For example, Tsukamurella species are invariably misidentified as Rhodococcus species by the API Coryne.15,48 Once a cli­nical isolate has been recognized as a Tsukamurella, standard phenotypic methods have limited usefulness for species-level identification, and the interpretation of these tests may be problematic for a number of reasons: only one or few strains have been included in the original species descriptions and the few subsequent articles; studies describing phenotypic properties of novel species have used predominantly home-brew methods and, occasionally, commercial media of different basic composition; recent characterizations of novel species have only included a limited set of phenotypic markers; there is conflicting information about certain differential physiological and biochemical characteristics of the various species; different media, whether commercial or home-brew, may yield inconsistent results, with either positive or negative reactions for the same phenotypic marker15; some biochemical reactions are variable among strains within the same species and even among different isolates of the same strain.28 Tsukamurella species are able to utilize a number of substrates including sugars as a sole carbon source and amino acids as a sole carbon and nitrogen source. Tsukamurella paurometabola is the most biochemically inert, whereas T. tyrosinosolvens, T. inchonensis, and T. carboxydivorans are the most active. In contrast to other Tsukamurellae, T.  carboxydivorans possesses a carbon monoxide dehydrogenase enzyme (CO-DH), which can be upregulated, and which enables it to grow under a CO-rich atmosphere of up to 300,000 p.p.m. T. tyrosinosolvens hydrolyzes tyrosine, a property shared by T. carboxydivorans. Other differential phenotypic properties are summarized in Table 18.1. Mycolic acids are very long-chain branched α-alkyl-βhydroxy fatty acids and the most characteristic component in the cell-wall lipids of aerobic actinomycetes that include Tsukamurellae. Tsukamurella species possess mycolic acids generally ranging in length from 62 to 78 carbon atoms with 1–7 double bonds.11 T. carboxydivorans contains longer mycolic acids with 81–95 carbons. Whole cell acid and alkaline methanolysates produce two adjacent mycolic acid spots, designated α and α', on thin-layer chromatography, which are unique to the genus Tsukamurellae.11–14 Reverse-phase highperformance liquid chromatographic analysis (HPLC) of mycolic acid esters yields a single cluster of poorly resolved peaks with retention times of 6.25–7.50 min.15 This profile

Molecular Detection of Human Bacterial Pathogens

is distinctive for members of the genus Tsukamurellae and allows their separation from other genera among the aerobic actinomycetes.11,15 However, this profile is fairly similar among Tsukamurella species; thus, HPLC analysis is not suitable for species-level identification.15 Quantitative analyses of pyrolysis products of mycolic acid methyl esters by gas chromatography and mass spectrometry using a standardized procedure on the Microbial Identification System (MIS Inc., Del.) yield straight-chain saturated and monounsaturated fatty acids ranging from 12 to 20 carbons in length and frequently tuberculostearic acid.15,20,21 The most abundant fatty acids are C16:0 and C18:1ω9c, which account for 65%–75% of fatty acid content in all species. Using the MIS library and pattern-recognition software, the shortchain fatty acid profiles of Tsukamurella isolates overlapped with that of other members of the aerobic actinomycetes, such as Nocardia, Rhodococcus, or Gordonia species with high similarity indices15,34; thus, MIS appeared not to be useful for genus-level identification. However, using a heuristic library approach developed with the MIS software from profiles of a large number of reference and clinical strains of aerobic actinomycetes, McNabb et al. were able to correctly assign eight clinical isolates to the genus Tsukamurella and to determine that they were closely related to T. inchonensis strain ATCC 25983 but not to T. paurometabola strain ATCC 8368.58 18.1.3.2  Molecular Techniques Novel genomic species of Tsukamurella have been characterized using DNA–DNA hybridization studies and 16S rRNA gene sequencing. The minimum level of DNA–DNA relatedness between strains required to delineate a novel species is 70%.59 Original species descriptions that included more than one strain invariably showed levels of DNA–DNA reassociation ≥70% among strains of the same species and positive hybridization, albeit at levels below this cutoff, among strains of different species. DNA–DNA hybridization ­methods, although a standard for taxonomists, are typically not performed in the clinical laboratory. The phylogenetic ­relationships among well-characterized reference and type strains of Tsukamurellae based on 16S rRNA sequences are shown in Figure 18.1. The 16S rRNA sequences of Tsukamurella species are closely related, with similarity levels exceeding 99% among most species. The lowest similarity index of 98.8% is observed between T. spongiae type strain and T. strandjordii, T. spumae, and T. pseudospumae. Tsukamurella ­sunchonensis and T. pseudospumae share an identical 16S rRNA sequence and very likely represent the same species. The most closely related species are T. tyrisonosolvens and T. carboxydivorans, with a 16S similarity index of 99.8%, followed by T. spumae and T. pseudospumae at 99.7%. Although the 16S sequences of isolates of the same species are usually identical, limited intraspecific polymorphisms have been observed. For example, in the original description by Yassin et al., one isolate of T. tyrosinosolvens showed only a 99.7% sequence similarity to the three other strains, includ­ ing the type strain, which had identical sequences. Similarly,

(+) (+) (−) (−) (−) (−)

Hydrolysis Esculin Urea Casein Hypoxanthine Xanthine Tyrosine (+) (+) (+) (+) (+) (+) (+) (−) (−) (+) (−) (+)

(+) (+) (−) (+) (−) (−)

(−) (+) (+) (−)

T. pulmonis

Notes:  (+): Positive; (−): Negative; NA: Not available.

Sugar utilization as sole carbon source d-Arabinose (−) l-Arabinose (−) d-Cellobiose (−) Dulcitol (−) meso-Erythritol (−) d-Fructose (+) d-Maltose (−) d-Mannitol (+) d-Melezitose (−) d-Ribose (+) d-Sorbitol (−) d-Xylose (+)

(+) (+) (+) (−)

Growth at 10°C 25–28°C 35–37°C 42–45°C

T. paurometabola

(+) (−) (−) (−) (−) (−) (+) (+) (+) (+) (+) (+)

(+) (+) (−) (+) (−) (−)

(−) (+) (+) (+)

T. inchonensis

(+) (+) (+) (+) (+) (+) (+) (+) (+) (+) (+) (+)

(+) (−) (−) (+) (−) (+)

(−) (+) (+) (−)

T. tyrosinosolvens

TABLE 18.1 Selected Differential Phenotypic Properties of Tsukamurella Species

(−) (−) (−) (−) (−) (−) (−) (+) (−) (+) (+) (−)

(−) (+) (−) (−) (−) (−)

(−) (+) (+) (−)

T. strandjordii

(−) NA NA NA (−) (+) (+) NA (+) NA (−) (−)

(+) (−) (−) (+) (−) (+)

(−) (+) (+) (−)

T. sunchonensis

(−) (+) (−) (+) (+) (+) (+) (+) (+) (+) (+) (+)

(−) (−) NA (+) NA (+)

(−) (+) (+) (−)

T. spumae

(+) (+) (−) (−) (−) (+) (+) (−) (+) (+) (−) (−)

(−) (−) NA (+) NA (+)

(+) (+) (+) (−)

T. pseudospumae

(−) (+) (+) (−) (−) (+) (−) (+) (+) (+) (+) (+)

NA NA NA NA NA NA

(−) (+) (+) (−)

T. spongiae

(+) (+) (+) (+) (+) (+) (+) (+) (+) (+) (+) (+)

(−) (+) (+) (+) (+) (+)

(−) (+) (+) (−)

T. carboxydivorans

Tsukamurella 193

194

Molecular Detection of Human Bacterial Pathogens

T. spongiae-AY714239-K362T T. spongiae-AY714240-K363 T. pulmonis-X92981-D1321T T. tyrosinosolvens-Y12248-D1411 908

1000

843

T. tyrosinosolvens-Y12246-D1397T T. carboxydivorans-EU521689-Y2T T. tyrosinosolvens-Y12245-D1498

266

981 T. inchonensis-AF283282-ATCC25938 452

T. inchonensis-AF283281-ATCC700082T T. paurometabola-AF283280-DSM20162T

419

T. strandjordii-AF283283-ATCCBAA173T

789

989

540

T. pseudospumae-AY238513-N1176T T. sunchonensis-AF150494-SCNU5T T. pseudospumae-AY333425-JC85 T. spumae-Z37150-N1171T 992 T. spumae-AY238512-N1173

0.0005

FIGURE 18.1  Neighbor-joining phylogenetic tree based on 16S rRNA sequences of reference and type strains of Tsukamurella species. Numbers at the nodes indicate bootstrap values. The scale at the bottom indicates the genetic distance. Species names are followed by GenBank accession numbers and strain names. “T” indicates the type strains.

the  published sequences of the two strains of T. spongiae have only 98.9% sequence similarity (Figure 18.1). In one study, partial sequencing of the first 500 bp of the 16S rRNA gene and comparison with sequences in a commercial database and in GenBank allowed genus-level identification of Tsukamurellae but species identification could only be achieved for 62% of isolates.60 However, alignment of fulllength 16S rRNA sequences of well-characterized strains of Tsukamurella disclosed species-specific signature sequences that could potentially be used for definitive species-level identification of unknown isolates (Table 18.2). Ribotyping has revealed that Tsukamurellae species may harbor up to four ribosomal operons.17 The sequences of other housekeeping genes such hsp65, rpoB, gyrB, groEL, and 16S-23S internal transcribed spacer were demonstrated to be phylogenetically informative and more discriminatory for species definition of other aerobic actinomycete genera, such as Mycobacterium and Nocardia. However, the number of published sequences for these genes is insufficient to draw similar conclusions regarding Tsukamurella species. The published groEL sequences of T.  paurometabola (AF352578) and T. tyrosinosolvens (U90204) show a difference of 5.4%. Restriction fragment length polymorphism (RFLP) analysis of hsp65 has shown that this method is reliable in distinguishing Tsukamurella paurometabola from other aerobic actinomycetes.61 The published gyrB sequences of T. paurometabola and T. inchonensis display a difference of 7.6%. These results indicate that building a library of more polymorphic sequences of

h­ ousekeeping genes could potentially improve the confidence level for identifying Tsukamurella species.

18.2  METHODS 16S rRNA gene sequencing for identification of Tsukamurella species from bacterial cultures is detailed below.

18.2.1  Sample Preparation Bacteria grown on any media are suitable for the test. Depending on the size of the colonies, 1–20 colonies from a pure culture are scraped off the agar with a disposable or flamed wire loop without disrupting the agar surface. Agar media contain substances that may inhibit the PCR reaction. Colonies of any age can be used, although freshly subcultured colonies are preferred. Bacteria can also be collected from a broth culture (e.g., blood culture bottle, trypticase soy broth, or brain heart infusion broth). About 1.5 ml of broth is collected with a sterile pipette in a 2 mL tube. Heat the tube at 95°C for 15 min in a dry heat block to inactivate the bacteria. Spin at 13,000 rpm for 5  min. The supernatant is discarded, leaving the bacterial pellet. The pellet is washed twice in 500  µL of 1× sterile Phosphate Buffered Saline (PBS) before proceeding to DNA extraction. The washing step is optional for bacteria collected from the surface of solid agar media. The pellet size should ideally not exceed 3 mm3. If the samples are not to be processed the same day, cell pellets can be stored frozen at –70°C.

T T T T T C T G C C A A A C C G A C G C A G T G

T T C G C C T G C A A A A C C G A G T C A G T G

C T C G C C C G C A G T A C C G A G T C A G C G

T T C G C C T G C C G T G C C G A C G T T G T G

T T C G C C T/C G T/C A/G G T A C C G A G T C A G T G

T T C G C C T G T A G T A C C G G G T C A G T A

T C Del Del C T C T C A G T A C C G A C G T T G T G

T C Del Del C T C T C A G T A C C G A G T C A A T G

T C Del Del C T C T C A G T A C C G A G T C A A T G

C T C G C C C G C A G T A T T C A G T C A G C G

T. paurometabola T. strandjordii T. pulmonis T. inchonensis T. tyrosinosolvens T. carboxydivorans T. spumae T. pseudospumae T. sunchonensis T. spongiae

Notes:  Positions are based on E. coli numbering (Ref. 65: Brosius et al. 1978). Del:  Deletion

86 88 89 90 145 190 191 203 215 381 546 560 595 737 739 743 923 1011 1018 1019 1136 1260 1267 1365

Position

TABLE 18.2 16S rRNA Gene Sequence Signatures That Differentiate Tsukamurella Species

Tsukamurella 195

196

Molecular Detection of Human Bacterial Pathogens

DNA is extracted with the Qiagen QIAamp DNA Mini Kit as follows: Add 180 µL of Buffer ATL (Lysis Buffer from the Qia­ gen QIAamp DNA Mini Kit) and 20 µL Proteinase K to the bacterial pellet and vortex vigorously. Incubate overnight at 56°C in a dry heat block. Proceed with the extraction using the tissue protocol of QIAamp DNA Mini Kit as described by the manufacturer.

18.2.2  Detection Procedures (i) PCR Amplification  The PCR mix consists of: HotStart Taq Polymerase

0.5 µL

10× PCR buffer

10 µL

P1*: 50 pmoles (Stock 25 µM)

2 µL

P10*: 50 pmoles (Stock 25 µM)

2 µL

dNTP mixture (25 mM each)

0.8 µL

MgCl2 (25 mM stock)

12 µL

Molecular biology grade ddH2O

62.7 µL

Bacterial DNA solution

10 µL

Total reaction volume

100 µL

The PCR and sequencing primers are:

1. P1* = 8FPL (F: 8–27): 5', AG AGT TTG ATC CTG GCT CAG 3', 2. P2 = 516F (F: 516–533): 5', T GCC AGC AGC CGC GGT AA 3', 3. P3 = 516R (R: 516–533): 5', TT ACC GCG GCT GCT GGC A 3', 4. P4 = 806F (F: 787–806): 5', ATT AGA TAC CCT GGT AGT CC 3', 5. P5 = 806R (R: 787–806): 5', GG ACT ACC AGG GTA TCT AAT 3', 6. P6 = 1175F (F: 1175–1194): 5', GAG GAA GGT GGG GAT GAC GT 7. P7 = 1175R (R: 1175–1194): 5', AC GTC ATC CCC ACC TTC CTC 3', 8. P10* = DG74 (R: 1522–1540): 5', A GGA GGT GAT CCA RCC GCA 3'.

(*) indicates PCR primers. Numbers in parentheses correspond to E. coli numbering positions.62 Primer sequences are derived from references.63–65 F = Forward; R = Reverse. For primer P10, R represents A or G. Each run includes a negative control, 10 µL of water added to the PCR mix instead of DNA, and a positive control, 10 µL of DNA prepared from a laboratory strain of E. coli. The thermocycling parameters are as follows: initial denaturation at 95°C for 10 min; followed by 30 cycles of 95°C for 30 s; 68°C for 30 s and 72°C for 45 s; and a final 72°C for 10 min.

If the PCR product is to be used the same day, it can be stored at 4°C or on ice; otherwise, storage at −20°C is recommended. (ii) DNA Gel Electrophoresis  Prepare 1% agarose gel by mixing the agarose powder with 1× Tris/Borate EDTA (TBE). Dissolve by microwaving on high until the solution is clear. Add 5 µL of Ethidium Bromide (10 mg/mL) per 100 mL of agarose solution. Add 10 µL of PCR product to each well, mixed, with 10 µL of 2× loading dye. Add 1× TBE to the electrophoresis chamber to cover the gel surface and run for 45 min at 80 V with a DNA ladder of the appropriate size. If a product of expected size is visualized (approximately 1500 bp), proceed to PCR product purification and sequencing. (iii) PCR Product Purification  Purify the PCR product with Qiagen MinElute PCR Purification Kit or equivalent PCR purification kit. Follow instructions in package insert. (iv) Standard Cycle Sequencing  Eight sequencing reactions are prepared for each sample with primers P1 to P7 and P10 (primer sequences are detailed above). The reaction final volume is 10 µL and consists of 4 µL of purified PCR product solution, 4 µL of Big Dye Ready Reaction mix (Applied Biosystems) and 2 µL of primer diluted to 1 μM (final concentration = 0.2 μM). The sequencing parameters are as follows: 1 cycle at 96°C for 1 min, followed by 25 cycles of 96°C for 10 s, 50°C for 5 s, and 60°C for 4 min. The sequencing reactions are purified using the Qiagen DyeEx Spin Kit or equivalent. Reactions are analyzed by capillary electrophoresis on an ABI 3130 Genetic Analyzer (Applied Biosystems) or equivalent. (v) Sequence Analysis  The sequencing chromatogram contigs are assembled and edited using sequencing analysis software, for example, ChromasPro (Technelysium Pty Ltd., Australia) in order to resolve ambiguous positions. The sequence of an unknown isolate is saved in FASTA format and searched against GenBank using BLAST (http:// www.ncbi.nlm.nih.gov/BLAST). If the unknown sequence matches that of a Tsukamurella species, place it in a single file together with 16S rRNA sequences of reference and type strains of Tsukamurella species. The GenBank accession numbers of these sequences are shown in Figure  18.1. Align these sequences using the CLUSTAL algorithm with CLUSTAL_X,66 BioEdit,67 (http://www.mbio.ncsu.edu/ BioEdit/BioEdit.html), or MEGA software.68 These are freely available on the world-wide Web. Perform phylogenetic analysis with the neighbor-joining algorithm for the purpose of species identification. Phylogenetic trees can be constructed and viewed with either MEGA (www.megasoftware.net) or PHYLIP (http://­evolution.genetics.washington.edu/phylip. html). Phylogenetic trees generated in CLUSTAL_X using the neighbor-joining algorithm can be viewed using either MEGA or the Treeview software (http://taxonomy.zoology.gla.ac.uk/rod/treeview.html). Alternatively, align the

197

Tsukamurella

Tsukamurella sequences with the E. Coli 16S rRNA sequence J01695.62 The polymorphic 16S rDNA sequence signatures as outlined in Table 18.2 may then be used to derive the species identification.

18.3  C  ONCLUSIONS AND FUTURE PERSPECTIVES Tsukamurella species are soil organisms that have emerged over the past decade as a rare but significant cause of opportunistic infections associated with indwelling foreign medical devices in immunocompromised patients. Infections in immunocompetent patients have been sporadically reported and present mostly as a tuberculosis-like illness. Despite the fact that Tsukamurellae are multidrug-resistant organisms, the mechanisms of antimicrobial resistance have not been studied. It is uncertain whether they harbor plasmids. Conventional phenotypic methods allow genus-level presumptive identification at best. The small number of reported Tsukamurella strains that have been studied phenotypically and their physiological variability make it difficult to fit a given strain within a described species. There are no standardized molecular typing methods for Tsukamurella species. In a study investigating a pseudooutbreak that spanned a 30-month period at a hospital in South Carolina, the authors used ribotyping, which demonstrated that the 12 outbreak isolates had a similar profile, which was distinct from other sporadic nonoutbreakrelated isolates.23 Common methods for molecular typing of other bacterial species such as pulsed field gel electrophoresis (PFGE), multilocus sequence typing (MLST), and multilocus variable number tandem repeat analysis (MLVA) need to be developed for Tsukamurellae. Such progress is hampered by the paucity of sequence information in public databases. As of August 27, 2009, there were 105 nucleotide sequences of named Tsukamurella species, 90% of which are 16S rRNA sequences. Alignment of the 16S sequences of well-characterized strains enabled us to identify polymorphic signature sequences that could potentially serve as probe targets for the design of real-time PCR assays for rapid species-level identification (Table 18.2). The genome of Tsukamurella paurometabola type strain has been recently sequenced, but its annotation has not yet been completed. Out of 8862 published Tsukamurellae protein sequences in GenBank, 8848 belong to the Tsukamurella paurometabola type strain. Sequencing of a T. tyrosinosolvens isolate genome is currently in progress. As more sequence information becomes available, new methods based on comparative genomics can potentially be developed for more rapid and definitive species identification and molecular typing of Tsukamurellae in a clinical setting.





REFERENCES

1. Zhi, X.Y., Li, W.J., and Stackebrandt, E., An update of the structure and 16S rRNA gene sequence-based definition of higher ranks of the class Actinobacteria, with the proposal of two new suborders and four new families and emended descriptions of the existing higher taxa, Int. J. Syst. Evol. Microbiol., 59, 589, 2009.





2. Pavlov, D. et al., Potentially pathogenic features of heterotrophic plate count bacteria isolated from treated and untreated drinking water, Int. J. Food Microbiol., 92, 275, 2004. 3. Collins, M.D. et al., Tsukamurella gen. nov., harboring Corynebacterium paurometabolum and Rhodococcus aurantiacus, Int. J. Syst. Bacteriol., 38, 385, 1988. 4. Lechevalier, M.P., and Lechevalier, H.A., Chemical composition as a criterion in the classification of aerobic actinomycetes, Int. J. Syst. Bacteriol., 20, 435, 1970. 5. Steinhaus, E.A., A study of the bacteria associated with thirty species of insects, J. Bacteriol., 42, 757, 1941. 6. Jones, D., A numerical taxonomic study of coryneform and related bacteria, J. Gen. Microbiol., 87, 52, 1975. 7. Tsukamura, M., and Mizuno, S., A new species Gordona aurantiaca occurring in sputa of patients with pulmonary disease, Kekkaku, 46, 93, 1971. 8. Tsukamura, M., and Yano, I., Rhodococcus sputi sp. nov., nom. rev., and Rhodococcus aurantiacus sp. nov., nom. rev., Int. J. Syst. Bacteriol., 35, 364, 1985. 9. Collins, M.D., and Jones, D., Lipid composition of Corynebacterium paurometabolum, FEMS Microbiol. Lett., 13, 13, 1982. 10. Goodfellow, M. et al., Chemical and numerical taxonomy of strains received as Gordona aurantiaca, J. Gen. Microbiol., 109, 57, 1978. 11. Koerner, R.J., Goodfellow, M., and Jones, A.L., The genus Dietzia: A new home for some known and emerging opportunist pathogens, FEMS Immunol. Med. Microbiol., 55, 296, 2009. 12. Yassin, A.F. et al., Tsukamurella inchonensis sp. nov., Int. J. Syst. Bacteriol., 45, 522, 1995. 13. Yassin, A.F. et al., Tsukamurella pulmonis sp. nov., Int. J. Syst. Bacteriol., 46, 429, 1996. 14. Yassin, A.F. et al., Tsukamurella tyrosinosolvens sp. nov., Int. J. Syst. Bacteriol., 47, 607, 1997. 15. Kattar, M.M. et al., Tsukamurella strandjordae sp. nov., a proposed new species causing sepsis, J. Clin. Microbiol., 39, 1467, 2001. 16. Seoung, C.N. et al., Tsukamurella sunchonensis, sp. nov., a bacterium associated with foam in activated sludge, J. Microbiol., 41, 83, 2003. 17. Nam, S.W. et al., Tsukamurella spumae sp. nov., a novel actinomycete associated with foaming in activated sludge plants, Syst. Appl. Microbiol., 26, 367, 2003. 18. Nam, S.W. et al., Tsukamurella pseudospumae sp. nov., a novel actinomycete isolated from activated sludge foam, Int. J. Syst. Evol. Microbiol., 54, 1209, 2004. 19. Goodfellow, M. et al., Activated sludge foaming: The true extent of actinomycete diversity, Water Sci. Technol., 37, 511, 1998. 20. Olson, J.B. et al., Tsukamurella spongiae sp. nov., a novel actinomycete isolated from a deep-water marine sponge, Int. J. Syst. Evol. Microbiol., 57, 1478, 2007. 21. Park, S.W., et al., Tsukamurella carboxydivorans sp. nov., a carbon monoxide-oxidizing actinomycete, Int. J. Syst. Evol. Microbiol., 59, 1541, 2009. 22. Jones, R.S. et al., Persistent bacteremia due to Tsukamurella paurometabolum in a patient undergoing hemodialysis: Case report and review, Clin. Infect. Dis., 18, 830, 1994. 23. Auerbach, S.B. et al., Outbreak of pseudoinfection with Tsukamurella paurometabolum traced to laboratory contamination: Efficacy of joint epidemiological and laboratory investigation, Clin. Infect. Dis., 14, 1015, 1992. 24. Haft, R.F. et al., Pseudoinfection due to Tsukamurella paurometabolum, Clin. Infect. Dis., 15, 883, 1992.

198 25. Tsukamura, M., and Kawakami, K., Lung infection caused by Gordona aurantiaca (Rhodococcus aurantiacus), J. Clin. Microbiol., 16, 604, 1982. 26. Prinz, G. et al., Meningitis caused by Gordona aurantiaca (Rhodococcus aurantiacus), J. Clin. Microbiol., 22, 472, 1985. 27. Casella, P., Tommasi, A., and Tortorano, A.M., Peritonie da Gordona aurantiaca (Rhodococcus aurantiacus) in dialisi peritoneale ambulatoria continua, Microbiologia Medica, 2, 47, 1987. 28. Tsukamura, M. et al., Severe progressive subcutaneous abscesses and necrotizing tenosynovitis caused by Rhodococcus aurantiacus, J. Clin. Microbiol., 26, 201, 1988. 29. Osoagbaka, O.U., Evidence for the pathogenic role of Rhodococcus species in pulmonary diseases, J. Appl. Bacteriol., 66, 497, 1989. 30. Shapiro, C.L. et al., Tsukamurella paurometabolum: A novel pathogen causing catheter-related bacteremia in patients with cancer. Clin. Infect. Dis., 14, 200, 1992. 31. Lai, K.K., A cancer patient with central venous catheterrelated sepsis caused by Tsukamurella paurometabolum (Gordona aurantiaca), Clin. Infect. Dis., 17, 285, 1993. 32. Granel, F. et al., Cutaneous infection caused by Tsukamurella paurometabolum, Clin. Infect. Dis., 23, 839, 1996. 33. Rey, D. et al., Tsukamurella infections. Review of the literature apropos of a case. Pathol Biol (Paris), 45, 60, 1997. 34. Chong, Y. et al., Tsukamurella inchonensis bacteremia in a patient who ingested hydrochloric acid, Clin. Infect. Dis., 24, 1267, 1997. 35. Larkin, J.A. et al., Infection of a knee prosthesis with Tsukamurella species, South Med. J., 92, 831, 1999. 36. Maertens, J. et al., Catheter-related bacteremia due to Tsukamurella pulmonis, Clin. Microbiol. Infect., 4, 51, 1998. 37. Schwartz, M.A. et al., Central venous catheter-related bacteremia due to Tsukamurella species in the immunocompromised host: A case series and review of the literature, Clin. Infect. Dis., 35, e72, 2002. 38. Sheridan, E.A. et al., Tsukamurella tyrosinosolvens intravascular catheter infection identified using 16S ribosomal DNA sequencing, Clin. Infect. Dis., 36, e69, 2003. 39. Alcaide, M.L., Espinoza, L., and Abbo, L., Cavitary pneumonia secondary to Tsukamurella in an AIDS patient. First case and a review of the literature, J. Infect., 49, 17, 2004. 40. Elshibly, S. et al., Central line-related bacteraemia due to Tsukamurella tyrosinosolvens in a haematology patient, Ulster Med. J., 74, 43, 2005. 41. Almehmi, A. et al., Implantable cardioverter-defibrillator infection caused by Tsukamurella, W.V. Med. J., 100, 185, 2004. 42. Shaer, A.J., and Gadegbeku, C.A., Tsukamurella peritonitis associated with continuous ambulatory peritoneal dialysis, Clin. Nephrol., 56, 241, 2001. 43. Perez, V.A., Swigris, J., and Ruoss, S.J., Coexistence of primary adenocarcinoma of the lung and Tsukamurella infection: A case report and review of the literature, J. Med. Case Reports, 2, 207, 2008. 44. Maalouf, R. et al., First case report of community-acquired pneumonia due to Tsukamurella pulmonis, Ann. Intern. Med., 150, 147, 2009. 45. Shim, H.E. et al., A case of catheter-related bacteremia of Tsukamurella pulmonis, Korean J. Lab. Med., 29, 41, 2009. 46. Moore, J.E., Xu, J., and Millar, B.C., Analysis of 16S-23S intergenic spacer regions of the rRNA operons in Tsukamurella pulmonis, Br. J. Biomed. Sci., 63, 25, 2006. 47. Matsumoto, T. et al., Tsukamurella tyrosinosolvens cultured from sputum of a patient who received total gastrectomy for gastric cancer, Kekkaku, 81, 487, 2006.

Molecular Detection of Human Bacterial Pathogens 48. Bouza, E. et al., Tsukamurella: A cause of catheter-related bloodstream infections, Eur. J. Clin. Microbiol. Infect. Dis., 28, 203, 2009. 49. Sheng, W.H. et al., Brain abscess caused by Tsukamurella tyrosinosolvens in an immunocompetent patient, J. Clin. Microbiol., 47, 1602, 2009. 50. Woo, P.C. et al., First report of Tsukamurella keratitis: Association between T. tyrosinosolvens and T. pulmonis and ophthalmologic infections, J. Clin. Microbiol., 47, 1953, 2009. 51. Woo, P.C. et al., Tsukamurella conjunctivitis: A novel clinical syndrome, J. Clin. Microbiol., 41, 3368, 2003. 52. Clausen, C., and Wallis, C.K., Bacteremia caused by Tsukamurella species, Clin. Microbiol. Newsl., 16, 6, 1994. 53. Rey, D. et al., Tsukamurella and HIV infection, AIDS, 9, 1379, 1995. 54. NCCLS: M24-A. Susceptibility testing of mycobacteria, nocardiae, and other aerobic actinomycetes; Approved standard. Wayne, PA: NCCLS, 2003. 55. Vollbrecht, E. et al., Production and structure elucidation of diand oligosaccharide lipids (biosurfactants) from Tsukamurella sp. nov., Appl. Microbiol. Biotechnol., 50, 530, 1998. 56. Asano, M. et al., Reciprocal action of interferon-gamma and interleukin-4 promotes granulomatous inflammation induced by Rhodococcus aurantiacus in mice, Immunology, 88, 394, 1996. 57. Stanley, T. et al., The potential misidentification of Tsukamurella pulmonis as an atypical Mycobacterium species: A cautionary tale, J. Med. Microbiol., 55, 475, 2007. 58. McNabb, A. et al., Fatty acid characterization of rapidly growing pathogenic aerobic actinomycetes as a means of identification, J. Clin. Microbiol., 35, 1361, 1997. 59. Wayne, L.G., Brenner, D.J., and Colwell, R.R., International committee on systematic bacteriology report of the ad hoc committee on reconciliation of approaches to bacterial systematics, Int. J. Syst. Bacteriol., 37, 463, 1987. 60. Patel, J.B. et al., Sequence-based identification of aerobic actinomycetes, J. Clin. Microbiol., 42, 2530, 2004. 61. Steingrube, V.A. et al., Rapid identification of clinically significant species and taxa of aerobic actinomycetes, including Actinomadura, Gordona, Nocardia, Rhodococcus, Streptomyces, and Tsukamurella isolates, by DNA amplification and restriction endonuclease analysis, J. Clin. Microbiol., 35, 817, 1997. 62. Brosius, J. et al., Complete nucleotide sequence of a 16S ribosomal RNA gene from Escherichia coli, Proc Natl Acad Sci USA, 75, 4801, 1978. 63. Fredericks, D.N., and Relman, D.A., Sequence-based identification of microbial pathogens: A reconsideration of Koch’s postulates, Clin. Microbiol. Rev., 9, 18, 1996. 64. Fredricks, D.N., and Relman, D.A., Improved amplification of microbial DNA from blood cultures by removal of the PCR inhibitor sodium polyanetholesulfonate, J. Clin. Microbiol., 36, 2810, 1998. 65. Greisen, K. et al., PCR primers and probes for the 16S rRNA gene of most species of pathogenic bacteria, including bacteria found in cerebrospinal fluid, J. Clin. Microbiol., 32, 335, 1994. 66. Thompson, J.D. et al., The CLUSTAL_X windows interface: Flexible strategies for multiple sequence alignment aided by quality analysis tools, Nucleic Acids Res., 25, 4876, 1997. 67. Hall, T.A., BioEdit: A user-friendly biological sequence alignment editor and analysis program for Windows 95/98/ NT, Nucleic Acids Symposium Series, 41, 95, 1999. 68. Tamura, K. et al., MEGA4: Molecular Evolutionary Genetics Analysis (MEGA) software version 4.0. Mol. Biol. Evol., 24, 1596, 2007.

Section II Firmicutes and Tenericutes Bacilli

19 Abiotrophia Marco Arosio and Annibale Raglio CONTENTS 19.1 Introduction...................................................................................................................................................................... 201 19.1.1 Classification, Morphology, and Biology............................................................................................................. 201 19.1.2 Clinical Features................................................................................................................................................... 202 19.1.3 Diagnosis.............................................................................................................................................................. 203 19.1.4 Antibiotic Susceptibility....................................................................................................................................... 206 19.2 Methods............................................................................................................................................................................ 208 19.2.1 Sample Preparation............................................................................................................................................... 208 19.2.2 Detection Procedures............................................................................................................................................ 208 19.3 Conclusions and Future Perspectives............................................................................................................................... 208 References.................................................................................................................................................................................. 209

19.1  INTRODUCTION The genus Abiotrophia, nowadays, is well known, but its definition has undergone extensive change. Only the application of molecular biology has made it possible to verify the correct classification of genus and species. The word “Abiotrophia” means “life nutrition deficiency” and refers to the species’ requirements for supplemented media for growth.1 The genus Abiotrophia was proposed to include microorganisms initially known as nutritionally variant streptococci (NVS), also called “satelliting streptococci,” which are fastidious bacteria that need additional growth factors to survive in complex media: l-cysteine or pyridoxal hydrochloride (vitamin B6) has been shown to favor growth.2 Frenkel and Hirsch described first Abiotrophia as ungroupable viridans streptococci that grew as satellite colonies around other bacteria.3 Because of sugar fermentation and cell wall composition, these bacteria appeared to be mutant subspecies of Streptococcus mitis, but later, important differences in the enzymatic properties and patterns of PBPs between NVS and S. mitis appeared.4 Many synonyms were given to these fastidious microorganisms, such as satelliting Streptococcus, thiol-requiring Streptococcus, vitamin B6–dependant Streptococcus, piridoxal-dependant Streptococcus, symbiotic Streptococcus, and nutritionally variant streptococci, and only later were they classified as a new genus, Abiotrophia.5 Following the study by Collins and Lawson, Abiotrophia defectiva has remained the only Abiotrophia species, and a new genus was proposed as Granulicatella for the species found in human beings: Granulicatella adiacens,

Granulicatella para-adiacens and Granulicatella elegans, and a species Granulicatella balaenopterae originating from a whale.6 A. defectiva is present in a normal human oral cavity, urogenital tract, and intestinal flora.7 Like other viridans group streptococci A. defectiva causes bacteremia and sepsis and is also an etiological agent of endocarditis and other infectious diseases.8–13 The diagnosis is difficult but not impossible since A. defectiva identification starts from the suspect to its isolation. We will describe the traditional techniques for A. defectiva isolation and identification and underline the application of molecular biology to confirm the identification of a cultivated strain or whether it is present in sterile body fluid.

19.1.1  Classification, Morphology, and Biology Classification. The so-called NVS were classified in different genera (Abiotrophia and Granulicatella) and were differentiated in five species by molecular biology. At the beginning, Bouvet et al. considered these microorganisms to be members of two novel streptococcal species, and so they called them Streptococcus defectivus and Streptococcus adjacens and showed the presence of two different species within the NVS by chromosomal DNA–DNA hybridisation.14 Bouvet’s hybridization studies revealed that S. defectivus and S. adjacens shared less than 10% DNA homology with each other and with other streptococcal species.6 The phylogenetic differentiation of these microorganisms from streptococci group, together with nutritional considerations, satellitism, and pyrrolidonyl arylamidase productions, led to the definition of the new genus Abiotrophia. 201

202

It was only after the application of molecular biology and the analysis of 16S rRNA sequences that Kawamura et al. proposed the creation, within genus Abiotrophia, of two new species A. defectiva and Abiotrophia adiacens.15 The Kawamura study using 16S rRNA sequence analysis allowed the exclusion of these NVS organisms from the genus Streptococcus based on phylogenetic features (e.g., 16S rRNA sequence is different from streptococcal species by more than 11%) and also whether they shared a higher 16S rRNA relatedness with other catalase-negative bacteria (e.g., Aerococcus, Carnobacterium, Enterococcus) than with streptococci.6 Later, two further species of genus Abiotrophia were characterized: Abiotrophia elegans from a patient with endocarditis and Abiotrophia balaenopterae from a whale (Balaenoptera acutorostrata).16,17 These new species presented a closer phylogenetic affinity with Abiotrophia adiacens (16S rRNA sequence differed about 3%) than with A. defectiva (sequence differed about 7%); therefore, two distinct lines were defined within the genus Abiotrophia, namely, A. defectiva and a group including Abiotrophia adiacens, Abiotrophia elegans, and Abiotrophia balaenopterae (not human).6 Kanamoto et al. observed the heterogeneity among Abiotrophia strains and proposed a fourth human pathogen, Abiotrophia para-adiacens.18 In the same year, on the basis of 16S rRNA studies, Collins and Lawson noted that the genus Abiotrophia could be restricted to species A. defective, which is phylogenetically closer to several non-Abiotrophia genera (Globicatella, Facklamia, and Eremococcus), while the new genus Granulicatella was proposed for the other four species Granulicatella adiacens, Granulicatella para-adiacens, Granulicatella elegans, and Granulicatella balaenopterae.6 Morphology. A. defectiva observation by electron microscope presents a normal streptococcal ultrastructure if the cells are cultured in appropriate conditions. However, if the conditions are not optimal, the cell wall will show abnormal features considered by some authors as L-forms organisms.1 Colonies of A. defectiva are small, measuring 0.2–0.5 mm in diameter, and are either nonhemolytic or α-hemolytic on supplemented blood agar culture. On Gram stain, cell may be pleomorphic and present variable staining features. The aspect of colonies as well as the cellular morphology to Gram stain could be very variable because of nutritional state of the microorganisms.4,19 Biology. A. defective, like all the NVS, is defined by its requirement of pyridoxal or thiol group supplementation for growth. It also presents the particular feature of forming satellite colonies, usually nonhemolytic or alpha-hemolytic, and a variety of gram-positive and gram-negative bacteria (staphylococci, streptococci, and family Enterobacteriaceae strains), as well as yeasts can support their growth.1 A. defectiva is able to grow in most blood culture media (with the notable exception of unsupplemented tryptic soy broth) because the pyridoxal present in human blood is sufficient to permit growth.4

Molecular Detection of Human Bacterial Pathogens

For subculture, however, solid media must be supple­ mented with 0.001% pyridoxal or 0.01% l-cysteine. Alternatively, the subculture plate can be cross-streaked with Staphylococcus aureus to provide these factors and support the growth of satellite colonies. Some studies suggest that blood cultures suspected of harboring NVS should be subcultured within 48 h because viability of the organism may decline with continued incubation.19 The original efforts to identify the factors required for growth of NVS focused on sulfhydryl-containing compounds. A lot of authors studied which supplement should have been the best. In particular, Frenkel and Hirsch reported that cysteine, thioglycolate, reduced glutathione, and thiomalic acid were able to help the growth of NVS.3 Other studies demonstrated that l-cysteine was a useful supplement for NVS growth. Cayeux et al. used brain heart infusion as a basal medium and obtained a good growth by either l-cysteine or glutathione in association with either thioglycolate or dithiothreitol.20 A number of authors showed that vitamin B6 analogs were able to help the growth of NVS. George found pyridoxine (vitamin B6) supplementation allowed for growth of NVS on horse blood agar.21 Carey and Robert found that pyridoxal and pyridoxamine (active forms of vitamin B6) were effective, but pyridoxine was inhibitory for an NVS strain they studied.1 These apparent discrepancies were resolved because the studies of Shiller and Roberts suggested that pyridoxine is not assimilated by NVS and can inhibit their growth in media supplemented with other forms of vitamin B6.1,22 Regarding biochemical and phenotypical classification Abio­ tophia shares the following characteristics: optochin resistance, vancomycin susceptibility, leucine aminopeptidase (LAP), l-pyrrolidonyl-beta-naphthylamide (PYR) and arylamidase production, negative reaction for gas production in De Man, Rogosa, and Share (MRS) broth, grows in 6.5% NaCl or at 10°C and 45°C, and no reaction on bile-esculin medium (Table 19.1). The genera Abiotrophia could be differentiated from Granulicatella by their patterns of carbohydrate fermentation and production of B-galactosidase. However, not all isolates of NVS can be easily classified at species level by biochemical and phenotypical tests. Only the application of molecular biology allows accurate species typing.

19.1.2  Clinical Features A. defectiva is a part of a normal flora of the oral cavity and the urogenital and intestinal tracts. Like other NVS normally resident in the oral cavity, Abiotrophia has been identified as agent of endocarditis involving both native and prosthetic valves. In addition to their involvement with endocarditis, Abiotrophia has been implicated in a variety of other infections. In particular, before the final typing of Abiotrophia defective, all the clinical cases were reported as caused by NVS.10–12,23–36

203

Abiotrophia

TABLE 19.1 Identification of Abiotrophia and Granulicatella Speciesa Species A. defectiva G. adiacens G. paraadiacens G. elegans

Argb

Hip

Pul

Suc

Tag

Tre

𝛂-Gal

𝛃-Glu

𝛃-Gal

Origin

– − − +

– – – v

+ – − −

+ + + +

v + − –

+ − − –

+ − − −

− v − −

+ − − −

human human human human

Source:  Adapted from: http://www.cdc.gov/ncidod/biotech/strep/strep-doc/section2.htm#table7 a All strains are positive for PYR and LAP and sensitive to vancomycin; negative reactions for gas production in MRS broth, growth in 6.5% NaCl or at 10 and 45°C. No reaction on bile-esculin medium. All strains require pyridoxal for growth and satillite around staphylococcus culture on blood agar plates. b Abbreviations: Arg, deamination of arginine; Hip, hyrolysis of hippurate; Pul, Suc, Tag, Tre, acid production from pullulan, sucrose, tagatose, & trehalose broth respectively; α-gal, production of α-galactosidase; β-glu, production of β-glucouronidase; β-gal, production of -β-galactosidase + = 85% or more of the strains positive, – = 15% or less than the strains positive, v = variable reactions (16 to 84% positive).

Reports documented the isolation of NVS from the blood of patients with postpartum or postabortal sepsis, cirrhosis, and pancreatic abscess.37 Isolation has also been made from patients with otitis media, otitis externa, wound infection, and vaginal discharge. NVS have been implicated in conjunctivitis and infectious crystalline keratopathy.3,21 In the recent literature, it is possible to find that A. defectiva has also been isolated in cases of ophthalmic infections (keratitis and postoperative endophthalmitis); a brain abscess following neurosurgery; peritonitis; osteomyelitis (multiple discitis, sacroiliitis); arthritis; sepsis in immunocompromised patients and iatrogenic meningitis following myelography; and spinal anesthesia.9,13,38–47 Endocarditis. Because Abiotrophia spp. are fastidious organisms, it is likely that most cases are misdiagnosed as culture-negative endocarditis; therefore, their role in endocarditis may be underestimated. Christensen et al. (2001) reported that G. adiacens and A.  defectiva accounted for 3% of bacteremia due to  α-hemolytic streptococcus-related species other than S. pneumoniae.48,49 Roberts underlined that NVS are the cause of 5%–6% endo­carditis, and these strains could justify a culture-negative result.50 Stein and Nelson showed that the mortality rate of NVS endocarditis was higher than that reported for endocarditis caused by other streptococci or enterococci. The NVS endocarditis was also more difficult to treat, and the authors proposed that this could be due to the slow growth of NVS.51 Probably, the prolonged L-phase that Abiotrophia seems to be able to live that has been reported by some authors, can explain this difficulties encountered in treatment as a mechanism of noninherited resistance. Other Site Infections. Due to the Abiotrophia growth characteristics, its role in different infection sites may be

underestimated, as for endocarditis. Only recently, since the development of enriched agar and broth media and improvements in molecular biology, it has been possible to detect some clinical cases in a more precise manner. In a recent report, corneal scrapings of four patients with severe infectious keratitis were analyzed by culture and PCR of 16S ribosomal RNA (rRNA), followed by direct sequen­ cing of the resulting amplicon. The medical history of the four patients included laser-assisted in situ keratomileusis, penetrating keratoplasty, and trauma. Only PCR was able to identify a possible pathogen in all four cases, while bacterial cultures were either negative or did not correspond to the clinical picture. The authors concluded that analysis of corneal scraping by 16S rRNA PCR should be considered as a supplement to standard microbiological procedures. However, the results of this relatively new method have to be interpreted carefully.52 Interestingly, A. defectiva and Gnanulicatella were identified by molecular methods from the same maxillary sinus. Recent results from a multicenter study corroborate that Abiotrophia and Granulicatella species are associated with chronic maxillary sinusitis and may contribute to antibiotic treatment failure.53 Other studies confirmed that molecular techniques can be useful to pinpoint Abiotrophia as a cause of brain abscess and meningitis.54,55

19.1.3  Diagnosis The diagnosis is not easy because A. defectiva is a fastidious microorganism, and it is likely that most cases are misdiagnosed as culture negative. As discussed above, the diagnosis can be done by conventional methods and/or by molecular biology. A. defectiva should also be suspected when Gram stains of biological specimens reveal microbial cells but are culture negative.

204

The following critical points are worth emphasizing: (i)  microbiologists should remember that A. defectiva cells may exhibit variable morphology and staining characteristics; (ii) supplementation of laboratory media is desirable and often necessary to encourage the growth of these organisms; (iii) once a suspected A. defectiva is cultured, its identity should be confirmed by establishing its requirement for pyridoxal or cysteine; (iv) final identification is possible using biochemical tests but molecular technique should be preferred because it is more accurate; and (v) molecular technique is also useful to identify the presence of A. defectiva in sterile body fluids.

FIGURE 19.1  Colonies of A. defectiva on chocolate agar with 5% horse blood after 18–24 hours.

FIGURE 19.2  Colonies of A. defectiva on chocolate agar with 5% horse blood after 36–48 hours.

Molecular Detection of Human Bacterial Pathogens

Conventional Techniques. A. defectiva is not able to grow on the majority of agar or broth media normally used in routine microbiology laboratory practice. In fact, A. defectiva can be isolated only by media that are supplemented with vitamin B6; pyridoxal hydrochloride (10–100 mg/L); or l-cysteine (100 mg/L) under an aerobic or anaerobic atmosphere. In particular, it does not grow on sheep blood agar or chocolate agar with trypticase soy or Mueller-Hinton or Todd-Hewitt, but it can be isolated by brain heart infusion, thioglycolate broth, Schaedler, Columbia and Brucella, or chocolate agar with 5% horse blood.56 The biochemical tests useful for the identification of A. defectiva are reported in Table 19.1. Tiny alpha-hemolytic or nonhemolytic colonies may appear after 18 h on the adequate solid media (Figures 19.1–19.3). Alternatively a Staphylococcus aureus strain is proposed by the Centers for Disease Control and Prevention (CDC) as helper to induce satellite growth (Figure 19.4, Table 19.2). Following improvement of the culture media by including cysteine, A. defectiva can be detected in routine blood cultures in 2 or 3 days. The microscopic examination reveals both morphological pleiomorphism and variable Gram staining (Figures 19.5 and 19.6). Conventional current methods of identification rely on enzymatic and physiological properties, using commercially available identification systems such as Rapid ID 32 STREPT (bioMerieux) or automatic systems. In literature, these methods are not so accurate and often do not discriminate the species level. It is underlined that better results can be obtained when a dense inoculum id used.57 Molecular Techniques. The development of molecular techniques was very helpful for the diagnosis of Abiotrophia infections.2 In particular, it is possible to identify Abiotrophia from its colonies or by looking for its DNA in clinical samples with negative cultures.

FIGURE 19.3  Colonies of A. defectiva on columbia agar with 5% horse blood after 66–72 hours.

Abiotrophia

205

FIGURE 19.5  Gram stain from colonies of A. defectiva on chocolate agar with 5% horse blood after 18–24 hours.

FIGURE 19.4  Satellite growth on blood agar after 18–24 hours.

TABLE 19.2 Satellite Test (SAT) I.

II. III.

IV.

V.

Principle Abiotrophia and Granulicatella are usually fastidious and will grow only on supplemented media or enriched chocolate agar. The purpose of the SAT test is to determine if the unknown streptococcal strain is a nutritionally variant streptococci (NVS). These strains require pyridoxal for growth. Growth of Staphylococcus aureus on a TSA sheep blood agar plate releases these factors allowing NVS to grow. An alternative to performing the SAT test is to test for the requirement of vitamin B6 (pyridoxal). Inoculum The cultures are typically received on chocolate agar slants. Reagents and Materials 1. Staphylococcus aureus ATCC-25923 2. TSA-sheep blood agar plate Procedure 1. Streak the submitted culture over the entire surface of a TSA-sheep blood agar plate with an inoculating loop. 2. A single streak of Staphylococcus aureus ATCC-25923 is then made across the middle of the agar plate. 3. Incubate the plate in a candle extinction jar or C02 incubator for 24 h or more. Reading and Interpretation If the strain is a NVS growth will appear only adjacent to the staphylococcus streak, about 2 to 5 mm wide. Streptococci that are not NVS will grow over the entire surface of the agar plate.

Source: Adapted from: http://www.cdc.gov/ncidod/biotech/strep/strep-doc/ section1.htm

FIGURE 19.6  Gram stain from colonies of A. defectiva on chocolate agar with 5% horse blood after 66–72 hours.

In the literature of 1990s, it is possible to confirm the identity of Abiotrophia by 16S rRNA gene PCR and restriction fragment length polymorphism analysis. This method was able to discriminate only A. defectiva from Granulicatella (formerly Abiotrophia) adjacens.58 Nowadays, the most commonly used technique is nucleotide-sequencing analysis of 16S rRNA and 740-bp rpoB gene fragment. In particular, 16S rRNA analysis is performed by two different approaches: a commercial method, MicroSeq 500–1500 (Applied Biosystems) and homemade tests using different primers59–61 (Table 19.3). The Microseq 500 16S rRNA–based bacterial identification system is useful for identification of most clinically important bacterial strains with ambiguous biochemical profiles. The PCR amplification of a 740-bp rpoB gene fragment followed by sequencing is a suitable molecular approach for the identification of Streptococcus, Enterococcus, Gemella, Abiotrophia, and Granulicatella isolates at the species level.62

206

Molecular Detection of Human Bacterial Pathogens

TABLE 19.3 Primers and Probes for PCR Detection of A. defectiva Primers for PCR (5'–3')

Primers for Sequencing (5'–3')

Target Gene

Reference

LPW200-F: GAGTTGCGAACGGGTGAG LPW205-R: CTTGTTACGACTTCACCC

LPW200-F: GAGTTGCGAACGGGTGAG LPW205-R: CTTGTTACGACTTCACCC LPW99-F: TTATTGGGCGTAAAGCGA LPW273-R: TTGCGGGACTTAACCCAAC

16S rRNA

PC-Y Woo et al.60

31-F: GCCTTAGGACCTGGTGGTTT 830-R: GTTGTAACCTTCCAWGTCAT

2333-F: AARYTIGGMCCTGAAGAAAT 3073-R: TGIARTTTRTCATCAACCATGTG

rpoB

M. Drancourt et al.62

F: AGAGTTTGATCCTGGCTCAG R: GGCTACCTTGTTACGACTT

Analysis by RFLP

16S rRNA

Ohara-Nemoto et al.58

F: TATTACCGCGGCTGCTGGCA R: TCAGATTGAACGCTGGCGGC

F: TATTACCGCGGCTGCTGGCA R: TCAGATTGAACGCTGGCGGC

16S rRNA

Mikkelsen et al.44

Bact-338-F: ACTCCTACGGGAGGCAGC Bact-1525-R: AAGGAGGTGATCCAGCC

M13-F: GTAAAACGACGGCCAGT M13-R: GGAAACAGCTATGACCATG

16S rRNA

Paju et al.53

13B-F: GTGAATACGTTCCCGGGCCT 6-R: GGGTTTYCCCCRTTCRGAAAT

13B-F: GTGAATACGTTCCCGGGCCT 6-R: GGGTTTYCCCCRTTCRGAAAT

16S rRNA

Tung et al.56

FIGURE 19.7  Public bank web page where is possible to enter the sequence.

A critical step in these techniques is the DNA extraction, particularly from sterile body fluids (blood, cerebrospinal fluid [CSF], and others). The sequence analysis can be performed by a public bank or using commercial software programs: the former has more data but is not always well controlled, while the latter present a low number of gene sequences of particular bacteria in the database (Figures 19.7 and 19.8).

19.1.4  Antibiotic Susceptibility Because of their growth requirements, uncommon isolation from human clinical specimens, and lack of standardized testing methodology and interpretation, there are limited antimicrobial susceptibility data for the fastidious micro­ organisms available.63 It often happens that the clinician asks for a susceptibility test, and the clinical microbiologist has trouble producing it.

Abiotrophia

207

FIGURE 19.8  Sequence of A. defectiva.

Fortunately, in 2006 the Clinical and Laboratory Standards Institute (CLSI) published new guidelines on antimicrobial susceptibility testing of infrequently encountered or fastidious bacteria in the document M45-A “Methods for Antimicrobial Dilution and Disk Susceptibility Testing of Infrequently Isolated or Fastidious Bacteria. Approved Guidelines.”64 The testing conditions reported by CLSI document are as follows:

The testing conditions reported by Etest application sheet EAS 008–2007 are as follows:

Medium: cation-adjusted Mueller-Hinton broth with lysed horse blood (19.5%–5% v/v) and 0.001% (i.e., 1 mcg/mL) pyridoxal hydrochloride. Inoculum: direct colony suspension, equivalent to a 0.5 Mc Farland standard. Incubation: 35°C, ambient air, 20–24 h.

The quality-control recommendation is to utilize Streptococcus pneumoniae ATCC 49619. A susceptibility test is usually not necessary for isolates from respiratory sources or wounds, while A. defectiva isolated from normally sterile sources may be justified in immunodeficient patients. A. defectiva may present decreased susceptibility to penicillin, resulting in greater difficulty in the treatment of patients with endocarditis. In an isolate from an immunosuppressed patient was reported fluoroquinolone resistance.64 For isolates of A. defective, a synergistic effect has been demonstrated between beta-lactam agents and aminoglycosides.65 Tuohy et al. observed the susceptibilities of all strains to clindamycin, rifampicin, levofloxacin, ofloxacin, and

The minimal quality-control recommendation is to utilize Streptococcus pneumoniae ATCC 49619. The agents to consider for primary testing are penicillin, vancomycin, and cefotaxime or ceftriaxone. It is also possible to utilize the Etest method (AB BIODISK) for the chemo-antibiotic susceptibility test, although in the United States it is for investigational use only.

Agar medium: chocolate Mueller-Hinton + 0.001% pyridoxal HCl + 0.01% cysteine or Isosensitest + 5% human blood + 0.001% pyridoxal Hcl + 0.01% cysteine Inoculum: suspension in broth to 1 McFarland Incubation: 35°C/5% CO2/20–24 h.

208

quinupristin-dalfopristin, and the percentage of isolates of A. defectiva examined that were susceptible to other agents tested was as follows: penicillin, 8%; amoxicillin, 92%; ceftriaxone, 83%; and meropenem, 100%.66 Zheng et al. referred high rates of beta lactam and macrolide resistance in a pediatric population of A. defectiva isolates.7 Because of the use of macrolides as alternatives in prophylaxis for infective endocarditis in patients allergic to penicillin, the observation of increasing macrolide resistance may lead to failure in an important proportion of these patients undergoing dental or other invasive procedures.

19.2  METHODS The conventional methods described in Section 19.1.3 are quite similar to those usually applied in a traditional microbiology laboratory. Also, the molecular biology methods applied for the diagnosis of Abiotrophia are the same ones usually proposed for the identification of a strain from a positive culture or for the research of the DNA of a strain in a broth culture or in a usually sterile body fluid. Here, we will describe the principal steps of the common molecular biology procedures.55

19.2.1  Sample Preparation Adequate Samples. (i) Colonies from a culture plate; (ii) positive broth or blood culture; (iii) sterile body fluid such as: CSF, samples from abscess, peritoneal, pericardial, and joint fluid; and (iv) biopsy such as corneal scrapings or native or prosthetic valves. DNA Extraction. Usually it is sufficient to select one colony from a culture plate or 1 mL of culture broth or 500 µL–1 mL of sterile body fluid. It is important to underline that in some cases the sterile body fluid extract should be diluted assuming the presence of inhibitors and in some others it is better to concentrate due to the risk of low DNA load. The extraction from colonies is easy and some authors propose a simple lysis procedure with suspension in distilled water heated at 100°C and centrifuged at 20.000 × g for 10 min. DNA preparation from body fluid, biopsy, or blood may be more time consuming. There are different types of extractions, some manual and some by automatic systems. There are a large number of companies in the market producing DNA-extraction kits and systems that perform well. The most commonly used manual methods are PrepMan Ultra Sample Preparation Reagent Protocol (Applied Biosystems) and QIAamp Mini Kit (Quiagen), and for both the procedure is very simple. An increasing number of automatic systems are available in the market due to the interest of the companies in this new diagnostic field.

19.2.2  Detection Procedures The amplification can be performed using both homemade and commercial methods. There are a lot homemade protocols. Among the commercial techniques, MicroSeq

Molecular Detection of Human Bacterial Pathogens

500/1500 16S rRNA Bacterial Identification PCR kit (Applied Biosystem) is widely used. DNA Extraction. QIAamp Mini Kit (QIAGEN) can be used for DNA extraction for all samples on duplicate. The total DNA must be eluted in 50 µL of AE buffer by modifying the procedure described by manufacturer (QIAGEN) and stored at −80°C until measurement in the PCR. Because some extracted DNAs do not amplify, it can be useful to dilute 1:3 supposing the presence of inhibitors. PCR Amplification and Sequencing. Using MicroSeq500  16S rRNA Bacterial Identification PCR kit (Applied Biosystems), 15 µL must be examined by amplification 500 bp to the 5' end of 16S rRNA gene. Amplification of the bacterial 16S rRNA is carried out in a GeneAmp 9600 thermal cycler (Applied Biosystems). Amplicon detection, although not mandatory, could be verified by 2% agarose gel electrophoresis previous purification with YM-100 Microconcentrators (Celera Diagnostics). The sequencing reactions could be performed with MicroSeq500 16S rRNA Bacterial Identification Sequencing Kit. Cycle reactions are purified by the use of Centri-Sep Spin Columns (Applied Biosystems), and the cycle-sequencing products are analyzed on 3130 Genetic Analyzer (Applied Biosystems) according to the manufacturer’s instructions. Data Analysis. Raw data are analyzed with the Sequencing Analysis software v. 5.2 (Applied Biosystems). Sequences from both strands were aligned by using NCBI BLAST 2 Sequence (http://www.ncbi.nlm.nih.gov/blast/ bl2seq/wblast2.cgi). The resulting consensus sequence, obtained from the double-stranded portion of the alignment, is aligned with sequences stored in GenBank (BLAST; http://www.ncbi.nlm.nih.gov/BLAST). Criteria for Identification. For designation to species or genus level, we propose using the identification criteria formulated by Drancourt et al.: the species level is defined as a 16S rRNA sequence similarity of ≥99% with the prototype strain sequence in GenBank; the genus level is defined as a similarity of ≥97% with the prototype strain sequence in GenBank; a score lower than 97% should not be considered acceptable. After amplification, there some methods available to identify the organism from which DNA is derived. Some authors proposed restriction fragment length polymorphism (RFLP) analysis, but the sequence analysis followed by comparison of the nucleotide sequence with a database is easy and widely used. There are public databases such as GeneBank or the EMBL, where the limit is the risk of sequence not well controlled.

19.3  C  ONCLUSIONS AND FUTURE PERSPECTIVES The microbiological history of A. defectiva is fascinating and at the same time very instructive. From the general definition of NVS in 1960s, we are able today to reach the correct diagnosis.

209

Abiotrophia

Since their original description in 1961, the NVS have evolved from laboratory curiosities into streptococcal species in 1990s, and in 2000, into well-defined genera (e.g., Abiotrophia, Granulicatella and Globicatella, Facklamia and Eremococcus) and species (A. defectiva, Granulicatella adiacens, Granulicatella para-adiacens, Granulicatella elegans, and Granulicatella balaenopterae). There are numerous reports of Abiotrophia as cause of endocarditis, and in recent years there has been an increasing number of cases related to different sites of infection: brain abscess, meningitidis, peritonitis, osteomyelitis, arthritis, otitis, and keratitis. It is now well recognized that attempts should be made to look for Abiotrophia in Gram stain–positive and culture-negative samples. The traditional microbiology of Abiotrophia is well defined, and the correct solid or liquid media are available. The satellite test is now proposed and standardized by CDC. The 2006 CLSI document M45-A published the guidelines for the antimicrobial susceptibility testing and the agents to consider. The diffusion of these indications will improve the knowledge about the real susceptibility of Abiotrophia and the clinical activity of penicillin and macrolides. The application of DNA amplification methods is useful for the correct classification of genera and species, and the sequence analysis of 16S rRNA allows identification of Abiotrophia as a cause of culture-negative endocarditis or other sites’ infections. Molecular biology opens new fields of study: this is true above all for the so-called fastidious or infrequently isolated bacteria such as Abiotrophia. At the moment, there are no studies on resistance genes by the application of molecular biology techniques in strains of Abiotrophia, but it is probable that we will be able to confirm their presence in the near future. Further studies will also provide insights into the mechanisms by which they produce disease or are able to live in the l-phase.



REFERENCES









1. Ruoff, K.L., Nutritionally variant streptococci, Clin. Microbiol. Rev., 4, 184, 1991. 2. Boutoille, D. et al., Abiotrophia related endocarditis: Contribution of molecular biology, Presse Med., 28, 2149, 1999. 3. Frenkel, A., and Hirsch, W., Spontaneous development of L forms of streptococci requiring secretion of other bacteria or sulphydryl compounds for normal growth, Nature, 191, 728, 1961. 4. Johson, C.C., and Tunkel, A.R., Viridans streptococci and groups C and G streptococci. In Mandell, G.L., Bennett, J.E., Dolin, R. (Editors), Principles and practice of Infectious Diseases, 4 ed., p. 1851, Churchill Livingstone, New York, 1995. 5. Brouqui, P., and Raoult, D., Endocarditis due to rare and fastidious bacteria, Clin. Microbiol. Rev., 14, 177, 2001. 6. Collins, M.D., and Lawson, P.A., The genus Abiotrophia (Kawamura et al.) is not monophyletic: Proposal of Granulicatella gen. nov., Granulicatella adiacens comb. nov., Granulicatella elegans comb. nov. and Granulicatella balaenopterae comb. nov., Int. J. Syst. Evol. Microbiol., 50, 365, 2000.













7. Zheng, X., Antimicrobial susceptibilities of invasive pediatric Abiotrophia and Granulicatella isolates, J. Clin. Microbiol., 42, 4323, 2004. 8. Layland, J. et al., Abiotrophia endocarditis: The great masquerader, Aust. N.Z. J. Psychiatry, 42, 991, 2008. 9. O’Connor, K.M., Williams, P., and Pergam, S.A., An unusual case of knee pain: Pseudogout and Abiotrophia defectiva infection, South. Med. J., 101, 961, 2008. 10. Kiernan, T.J., Abiotrophia defectiva endocarditis and associated hemophagocytic syndrome—a first case report and review of the literature, Int. J. Infect. Dis., 12, 478, 2008. 11. Yerebakan, C., Aortic valve endocarditis due to Abiotrophia defectiva: A rare etiology, Wien Med. Wochenschr., 158, 152, 2008. 12. Assche, A.F., and Stephens, D.P., Infective endocarditis with Abiotrophia defectiva: The first Australian experience, Crit. Care Resusc., 10, 54, 2008. 13. Taylor, C.E., and Fang, M.A., Septic arthritis caused by Abiotrophia defectiva, Arthritis Rheum., 55, 976, 2006. 14. Bouvet, A. et al., Streptococcus defectivus sp. nov. and Streptococcus adjacens sp. nov. Nutritionally variant streptococci from human clinical specimens, Int. J. Syst. Bacteriol., 39, 290, 1989. 15. Kawamura, Y. et al., Transfer of Streptococcus adjacens and Streptococcus defectivus to Abiotrophia gen. nov. as Abiotrophia adiacens comb. nov. and Abiotrophia defectiva comb. nov., respectively, Int. J. Syst. Bacteriol., 45, 798, 1995. 16. Lawson, P.A. et al., Abiotrophia balaenopterae sp. nov., isolated from the minke whale (Balaenoptera acutorostrata), Int. J. Syst. Bacteriol., 49, 503, 1999. 17. Roggenkamp, A. et al., Abiotrophia elegans sp. nov., a possible pathogen in patients with culture-negative endocarditis, J. Clin. Microbiol., 36, 100, 1998. 18. Kanamoto, T., Sato, S., and Inoue, M., Genetic heterogeneities and phenotypic characteristics of strains of the genus Abiotrophia and proposal of Abiotrophia para-adiacens sp. nov., J. Clin. Microbiol., 8, 492, 2000. 19. Gross, K.C. et al., Evaluation of blood culture media for isolation of pyridoxal-dependent Streptococcus mitis, J. Clin. Microbiol., 14, 266, 1981. 20. Cayeux, P. et al., Bacterial persistance in streptococcal endocarditis due thiol-requiring mutants, J. Infect. Dis., 124, 247, 1971. 21. George, R.H., The isolation of symbiotic streptococci, J. Med. Microbiol., 7, 77, 1974. 22. Schiller, N.L., and Roberts, R.B., Vitamin B6 requirements of nutritionally variant Streptococcus mitior (mitis), J. Clin. Microbiol., 15, 740, 1982. 23. Lopardo, H., Mastroianni, A., and Casimir, L., Bacteremia due to Abiotrophia defectiva in a febrile neutropenic pediatric patient, Rev. Argent. Microbiol., 39, 93, 2007. 24. Takayama, R. et al., A case of isolated tricuspid valve infective endocarditis caused by Abiotrophia defectiva, Int. J. Cardiol., 118, 3, 2007. 25. Senn, L., Entenza, J.M., and Prod’hom, G., Adherence of Abiotrophia defectiva and Granulicatella species to fibronectin: is there a link with endovascular infections? FEMS Immunol. Med. Microbiol., 48, 215, 2006. 26. Yemisen, M. et al., Abiotrophia defectiva: A rare cause of infective endocarditis, Scand. J. Infect. Dis., 38, 939, 2006. 27. Senn, L. et al., Bloodstream and endovascular infections due to Abiotrophia defectiva and Granulicatella species, BMC Infect. Dis., 6, 9, 2006.

210 28. Sharaf, M.A., and Shaikh, N., Abiotrophia endocarditis: Case report and review of the literature, Can. J. Cardiol., 21, 1309, 2005. 29. Lainscak, M. et al., Infective endocarditis due to Abiotrophia defectiva: A report of two cases, J. Heart Valve Dis., 14, 33, 2005. 30. Hashimoto, T. et al., A woman with infectious endocarditis caused by Abiotrophia defectiva, Intern. Med., 43, 1000, 2004. 31. Raff, G.W. et al., Aortitis in a child with Abiotrophia defectiva endocarditis, Pediatr. Infect. Dis. J., 23, 574, 2004. 32. Chang, H.H. et al., Endocarditis caused by Abiotrophia defectiva in children, Pediatr. Infect. Dis. J., 21, 697, 2002. 33. Ray, M. et al., Infective endocarditis in a child due to Abiotrophia defectivus, Indian Pediatr., 39, 388, 2002. 34. Wijetunga, M., Bello, E., and Schatz, I., Abiotrophia endocarditis: A case report and review of literature, Hawaii Med. J., 61, 10, 2002. 35. Leonard, M.K., Pox, C.P., and Stephens, D.S., Abiotrophia species bacteremia and a mycotic aneurysm in an intravenous drug abuser, N. Engl. J. Med., 344, 233, 2001. 36. Okada, Y. et al., Endocardiac infectivity and binding to extracellular matrix proteins of oral Abiotrophia species, FEMS Immunol. Med. Microbiol., 27, 257, 2000. 37. McCarthy, L.R., and Bottone, E.J., Bacteremia and endocarditis caused by satelliting streptococci, Am. J. Clin. Pathol., 61, 585, 1974. 38. Arslan, U. et al., First case of peritonitis due to Abiotrophia defectiva, Perit. Dial. Int., 26, 725, 2006. 39. Esteban, J. et al., Postoperative endophthalmitis due to Abiotrophia defectiva, Enferm. Infecc. Microbiol. Clin., 23, 455, 2005. 40. Wilhelm, N. et al., First case of multiple discitis and sacroiliitis due to Abiotrophia defectiva, Eur. J. Clin. Microbiol. Infect. Dis., 24, 76, 2005. 41. Zenone, T., and Durand, D.V., Brain abscesses caused by Abiotrophia defectiva: Complication of immunosuppressive therapy in a patient with connective-tissue disease, Scand. J. Infect. Dis., 36, 497, 2004. 42. Cerceo, E. et al., Central nervous system infections due to Abiotrophia and Granulicatella species: an emerging challenge? Diagn. Microbiol. Infect. Dis., 48, 161, 2004. 43. Ince, A. et al., Total knee arthroplasty infection due to Abiotrophia defectiva, J. Med. Microbiol., 51, 899, 2002. 44. Mikkelsen, L., Theilade, E., and Poulsen, K., Abiotrophia species in early dental plaque, Oral. Microbiol. Immunol., 15, 263, 2000. 45. Michelow, I.C. et al., Abiotrophia spp. brain abscess in a child with Down’s syndrome, Pediatr. Infect. Dis. J., 19, 760, 2000. 46. Namdari, H. et al., Abiotrophia species as a cause of endophthalmitis following cataract extraction, J. Clin. Microbiol., 37, 1564, 1999. 47. Schlegel, L. et al., Iatrogenic meningitis due to Abiotrophia defectiva after myelography, Clin. Infect. Dis., 28, 155, 1999. 48. Christensen, J.J., and Facklam, R.R., Granulicatella and Abiotrophia species from human clinical specimens, J. Clin. Microbiol., 39, 3520, 2001.

Molecular Detection of Human Bacterial Pathogens 49. Christensen, J.J., Gruhn, N., and Facklam, R.R., Endocarditis caused by Abiotrophia species, Scand. J. Infect. Dis., 31, 210, 1999. 50. Roberts, R.B. et al., Viridans streptococcal endocarditis: The role of various species, including pyridoxal-dependent streptococci, Rev. Infect. Dis., 1, 955, 1979. 51. Stein, D.S., and Nelson K.E., Endocarditis due to nutritionally deficient streptococci: Therapeutic dilemma, Rev. infect. Dis., 9, 908, 1987. 52. Thiemo, R. et al., 16S rDNA PCR analysis of infectious keratitis: A case series, Acta Ophthalmol. Scand., 82, 463, 2004. 53. Paju, S. et al., Molecular analysis of bacterial flora associated with chronically inflamed maxillary sinuses, J. Med. Microbiol., 52, 591, 2003. 54. Biermann, C. et al., Isolation of Abiotrophia adiacens from a brain abscess which developed in a patient after neurosurgery, J. Clin. Microbiol., 37, 769, 1999. 55. Arosio, M. et al., Evaluation of the MicroSeq 500 16 S rDNAbased gene sequencing for the diagnosis of culture-negative bacterial meningitis, New Microbiol., 31, 343, 2008. 56. Koneman, E. et al., Color Atlas and Textbook of Diagnostic Microbiology, 4th ed., p. 389, J.B. Lippincott Company, Philadelphia, 1992. 57. Ruoff, K.L., Aerococcus, Abiotrophia, and other aerobic catalase-negative, gram-positive cocci. In Murray P.R. et al. (Editors), Manual of Clinical Microbiology, 9th ed., vol. 1, p. 443, ASM press, Washington DC, 2007. 58. Ohara-Nemoto, Y. et al., Identification of Abiotrophia adiacens and Abiotrophia defectiva by 16S rRNA gene PCR and restriction fragment length polymorphism analysis, J. Clin. Microbiol., 35, 2458, 1997. 59. Tung, S.K. et al., Identification of species of Abiotrophia, Enterococcus, Granulicatella and Streptococcus by sequence analysis of the ribosomal 16S-23S intergenic spacer region, J. Med. Microbiol., 56, 504, 2007. 60. Woo, P.C. et al., Granulicatella adiacens and Abiotrophia defectiva bacteraemia characterized by 16S rRNA gene sequencing, J. Med. Microbiol., 52, 137, 2003. 61. Roggenkamp, A. et al., PCR for detection and identification of Abiotrophia spp., J. Clin. Microbiol., 36, 2844, 1998. 62. Drancourt, M. et al., rpoB gene sequence-based identification of aerobic Gram-positive cocci of the genera Streptococcus, Enterococcus, Gemella, Abiotrophia, and Granulicatella, J. Clin. Microbiol., 42, 497, 2004. 63. Jorgensen, J.H., and Hindler, J.F., New consensus guidelines from the clinical and laboratory standards institute for antimicrobial susceptibility testing of infrequently isolated or fastidious bacteria, Clin. Infect. Dis., 44, 280, 2007. 64. CLSI, Methods for antimicrobial dilution and disk susceptibility testing of infrequently isolated or fastidious bacteria; approved guideline. M45-A, 2006. 65. Murray, C.K. et al., Abiotrophia bacteremia in a patient with neutropenic fever and antimicrobial susceptibility testing of Abiotrophia isolates, Clin. Infect. Dis., 32, E140, 2001. 66. Tuohy, M.J., Procop, G.W., and Washington, J.A., Antimicro­ bial susceptibility of Abiotrophia adiacens and Abiotrophia defectiva, Diagn. Microbiol. Infect. Dis., 38, 189, 2000.

20 Aerococcus Debarati Paul, D. Madhusudan Reddy, and Dipalok Mukherjee CONTENTS 20.1 Introduction.......................................................................................................................................................................211 20.1.1 Classification, Morphology, and Biology..............................................................................................................211 20.1.2 Clinical Features and Pathogenesis...................................................................................................................... 212 20.1.3 Diagnosis.............................................................................................................................................................. 213 20.2 Methods............................................................................................................................................................................ 215 20.2.1 Sample Preparation............................................................................................................................................... 215 20.2.2 Detection Procedures............................................................................................................................................ 215 20.3 Conclusion.........................................................................................................................................................................216 Acknowledgments.......................................................................................................................................................................216 References...................................................................................................................................................................................216

20.1  INTRODUCTION 20.1.1  Classification, Morphology, and Biology Aerococcus is a genus of aerobic gram-positive cocci that resemble enterococci but do not form chains. They are frequently isolated as airborne saprophytes in hospitals and as a pathogen of lobsters, cause greening in blood agar, and grow in the presence of 40% bile. In humans, they are found in endocarditis and in urinary tract infections and also some other health-related problems such as spondylodiscitis, lymphadenitis, and so forth.1–3 The genus Aerococcus was initially described by Williams.4 Aerococcus viridans has been isolated from a broad range of habitats such as air, vegetation, dust, soil, and meat-curing brines, and has also been associated, although rarely, with human infections. Despite its presence in such diverse sources, the taxonomic kinship of strains has been consistently demonstrated.5 Later on, Aerococcus urinae was proposed as a new species based on analysis of 16S rRNA sequences.6 A. urinae was isolated from urine of elderly persons suffering from urinary tract infections and from blood of patients with endocarditis and urosepticemia. Aerococcus spp. is a rarely reported pathogen, possibly due to difficulties in the identification of the organism. A. urinae is a gram-positive coccus that grows in pairs and clusters, produces alpha-hemolysis on blood agar, and is negative for catalase and pyrrolidonyl aminopeptidase. A. viridans organisms are gram-positive cocci with a strong tendency to form tetrads. A. urinae and A. viridans slightly differ in their reactions in the LAP and pyronylaminopeptidase (PYR) disk tests. A. urinae has the characteristics of being LAP-positive and PYR-negative,

whereas A. viridans is LAP-negative and PYR-positive.7,8 More recently, three additional species of Aerococcus have been described, based on 16S rRNA sequences and polyphasic taxonomy.9 Aerococcus urinaehominis has been recovered from a urine culture,5 Aerococcus sanguicola has been isolated from urine and blood,10 and A. christensenii has been recovered from a vaginal specimen.11 In general, the genus Aerococcus consists of bacterial strains that are gram-positive, nonspore-forming cocci that occurred as single cells, in pairs, tetrads, or in small groups. The cocci are facultatively anaerobic and catalase- and oxidase-negative. When grown on Columbia horse blood agar for 24 h, small (<1 mm diameter), entire, and grey colonies appear. Cells are nonmotile, and a-hemolysis is observed on blood agar.12 A. urinae is found in a constant but low percentage of urine specimens (0 ± 8%), but nearly all patients present clinical signs of acute infection. Although A. viridans has been shown to be responsible for a number of clinical conditions, a large proportion of strains isolated from clinical samples fail to exhibit any clinical manifestations.5,13 A. urinaehominis, also originated from a human urine sample, but no further clinical details were available on the organism, and its pathogenic potential remains unknown. A. christensenii is another strain that was not reported to cause disease and would not be discussed further. The absence of some of the important distinguishing characteristic features of this genus from the databases of most commercial identification systems may allow A. urinae to be misidentified as a staphylococci, streptococci, enterococci, pediococci, lactococci, or leuconostocs. Bosley et al. conducted experiments to differentiate among these genera.14 211

212

The cellular fatty acid profile of aerococci is distinguishable from other genera; two relatively novel fatty acids found in the aerococci are identified as C16:1 omega 9c and C16:1 omega 9t. DNA–DNA reassociation studies using 11 strains of aerococci with the type strain A. viridans ATCC 11563 show that DNA from eight of these strains are more than 75% related to the type strain and had 1%–4% divergence in related sequences. The remaining three strains are 60%–70% related to the type strain, have 7%–11.5% divergence, and may represent a second species, Aerococcus genospecies 2, which may now be known as A. urinae. Beta-glucuronidase, alpha-galactosidase, and beta-galactosidase are useful in characterizing the aerococci.

20.1.2  Clinical Features and Pathogenesis Patients infected with Aerococcus urinae tend to show one or more symptoms of urinary tract infection: dysuria (50%), pollakisuria (60%), abdominal pain (32.5%), flank tenderness (10%), fever, and general discomfort. Most patients have local or systemic conditions predisposing to urinary tract infections. Men have significantly more local problems, such as prostrate or urethral hyperplasia. Women have more systemic problems, such as dementia or cerebrovascular accidents. Men are in better physical condition than women.15 A retrospective chart review is performed by SierraHoffman et al. on 54 patients with positive urine cultures during a 1-year period to assess the clinical significance of A. urinae.9 The patients are classified as either having a UTI (urinary tract infection) or being “colonized,” depending on clinical and laboratory data. Patients are considered to have a UTI if they have >5 urine white blood cells and ≥105 CFU/mL on urine culture. They also are required to have at least two clinical symptoms (or only one symptom if catheterized). Clinical symptoms included are dysuria, frequency, nocturia, fever, myalgias, chills, altered mental status, and suprapubic pain. Patients with A. urinae recovered, but not meeting these criteria are classified as being “colonized.” There is an overall female predominance (91% in the UTI group and 95% in the colonized group). However, there are no differences in the age, sex, and underlying conditions in patients whose isolates are judged to represent a UTI as opposed to being colonized, with the exception that more patients with a urinary catheter are noted in the UTI group (P < .01). Both groups have a significant but equal incidence of underlying urologic conditions. • A. sanguinicola is similar to A. urinae in clinical implication, as both are first described as causes of urinary tract infections and are later found to be associated with bacteremia, sepsis, and endocarditis. A. viridans, the first Aerococcus species described, has been isolated from patients with meningitis, endocarditis, arthritis,16 and bacteremia in neutropenia and from HIV patients. • A. viridans is rarely associated with human infections; however, it could be a potential causative agent

Molecular Detection of Human Bacterial Pathogens

of bacteremia, especially in immunocompromised patients. Jafar et al. reported the first case of A. viridans bacteremia in a patient with HIV infection.17 The presumed route of infection in this patient is esophageal ulcers. • Infections due to this organism presumably seem to occur in previously damaged tissues or may be nosocomial in association with a prolonged hospitalization, antibiotic treatment, invasive procedures, presence of foreign bodies, or a neutropenic state.18,19 Kern and Vanek20 suggested that both granulocytopenia and oral mucositis may be the major risk factors for aerococcal bacteremia. Swanson et al.21 also described that A. viridans might be a significant pathogen in patients with functional asplenia, in that the organism is encapsulated with an acidic polysaccharide, and the strains with heavier encapsulation are more virulent.22 Recently, Nasoodi et al.23 reported spondylodiscitis due to A. viridans infection. The patient had low-grade back pain. The authors hypothesized that this man was immunocompromised due to his liver cirrhosis and became bacteremic following inoculation from a fish hook injury or following invasion from his gastrointestinal system after eating infected imported crustaceans. Infection is common in lobsters, causing Gaffkaemia,19 and other crustaceans, such as crab and some prawn species.23 Apart from urinary tract infections, A. urinae also may be associated with severe infections, such as endocarditis24 and septicemia. Astudillo et al. reported the first case of A. urinae spondylodiscitis,2 followed by Tekin et al.24 In this case, severe back pain was the major condition that the patient paid attention to and sought for help for. Surprisingly, despite the fact that urinary tract is the main route of entry of the organism, urine culture was found to be sterile. Even though A.  urinae endocarditis is clinically indistinguishable from endocarditis caused by other bacterial agents, infective endocarditis due to A. urinae has a poor prognosis. Late diagnosis is a possibility because of the subacute character of the disease and the unspecific symptoms of the elderly with comorbidities.25 Santos et al. reported a case of lymphadenitis caused by A. urinae.3 As for other urinary tract pathogens, the gastrointestinal tract could be a natural reservoir for A. urinae, though this has not been studied. Urinary tract infection is almost always the focus for an evolving bacteremia. An 80-year-old male with preexisting coronary artery disease and dementia and was admitted to the hospital for altered mental status. The patient developed slowly worsening urinary symptoms, including nocturia and frequency, over the 3 weeks prior to admission. Urine cultures grew >105 CFU of A. urinae, the only isolate, per mL. His urinary symptoms resolved and his mental status improved by hospital day 3.26 In another case, a 58-year-old person suffered from several days of dysuria, increased urinary frequency, and nocturia. Urinalysis revealed no bacteria, trace amounts of leukocyte

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Aerococcus

esterase, and negative results for nitrites, protein, ketones, bilirubin, and blood. A second urinalysis demonstrated a moderate level of leukocyte esterase, and negative urine chemistries. The urine culture on the second urine sample later grew >105 CFU of A. urinae, the only isolate, per mL.26 The case studies indicate that A. urinae is a rarely reported pathogen, possibly due to difficulties in the identification of the organism.26 Several conventional and molecular methods are therefore used for detection of the pathogen. In a case of urinary tract infections caused by A. urinae, the patient was treated empirically with intravenous ciprofloxacin.26 Urine cultures grew >105 CFU of A. urinae, the only isolate, per mL. His urinary symptoms resolved and his mental status improved by hospital day 3. The patient was discharged after oral ciprofloxacin was prescribed. Results of repeated urinalysis after discharge were normal. A. viridans strains are naturally susceptible to penicillin, macrolides and related drugs, tetracyclines, and chloramphenicol, and they are intrinsically resistant at a low level to aminoglycosides.1 In a case of endocarditis caused by A.  viridans in a 10-year-old child, he received ampicillin (200 mg/kg/day) and gentamicin (7.5 mg/kg/day) besides decongestive measures. Since the patient’s condition deteriorated during hospital stay, amikacin (15 mg/kg/day) and norfloxacin (15 mg/kg/day) were started on sixth day based on antibiogram. Improvement was noticed from 11th day of hospital stay with a gradual return of fever to baseline. However, at this point the patient developed severe chest pain and a pericardial rub. Echocardiography confirmed the presence of a pericardial effusion. The pericardial fluid was tapped. It was found to be sterile on culture, and steroids were started. The effusion gradually disappeared over the next 5 days, and the cardiac failure was also controlled. Amikacin and norfloxacin were continued for a total duration of 3 weeks.27 These case studies indicate that the treatment is often case sensitive and also depends on the species and its antibiotic susceptibility. A. viridans was found to be resistant to at least one antibiotic amongst eight wild type strains: erythromycin (six strains), tetracycline and minocycline (five strains), chloramphenicol (one strain), and high levels of streptomycin (one strain).1 Augustine et al.27 reported a case of endocarditis caused by A. viridans with multidrug resistance, that is, resistance to penicillin, ampicillin, cefotaxime, gentamicin, and intermediate resistant to ciprofloxacin, but they did not discuss minimum inhibitory concentrations (MICs). According to the antimicrobial susceptibility data of 30 aerococcal isolates obtained from Centers for Disease Control and Prevention,14 the MICs for nine strains were 0.5 g or more of penicillin per mL, with MICs for five strains being more than 1 g/mL; therefore, approximately 46% of aerococci tested were either relatively resistant or resistant to penicillin. Moreover, Christensen et al.28 recently documented that penicillin resistance should be the peculiar characteristic of A. viridans capable of differentiating it from Aerococcus-like organisms.19 Certainly, further investigations are needed to establish the optimal treatment for this pathogen.

In a case of endocarditis caused by A. urinae, treatment modalities were available for 10/11 patients.25 Among nine of these patients treated with a combination of a beta-lactam and an aminoglycoside, seven died. The remaining patient was treated with a beta-lactam alone and survived with neurologic sequelae. In all lethal cases, death occurred during the first 4 weeks (mean, 12 days). It is suggested that all patients are treated intravenously for a minimum of 6 weeks, since all surviving patients, including the presented patient, were treated for this time period either with a beta-lactam alone or with a beta-lactam and an aminoglycoside.

20.1.3  Diagnosis The concept of polyphasic taxonomy integrates different kinds of data and information on microorganisms, that is, phenotypic, genotypic, and phylogenetic, and this method has gained increasing acceptance since its introduction.29 This method includes transmission electron microscopy, fatty acid methyl ester analysis, multilocus enzyme electrophoresis, antimicrobial susceptibility tests, and comparison of 16S rRNA and/or other housekeeping gene(s) sequence data, and ribotyping. After the discovery of A. viridans, all the other strains were discovered on the basis of polyphasic taxonomy. Colony Morphology, Staining, and Biochemical Tests. The presumptive identification of aerobic gram-positive alpha-hemolytic cocci and the decision on whether to more fully identify the isolates are frequently based on Gram stain, colony appearance, and catalase reaction. Because A. urinae is catalase-negative, it could be mistaken for alpha-hemolytic streptococci or enterococci, which are more common urine isolates. At 24 h, the colony morphology resembles that of an alpha-hemolytic streptococcus or lactobacillus; at 48 h, it is similar to that of an Enterococcus. The Gram stain should be differential because A. urinae forms pairs, tetrads, and clusters. However, since A. urinae shows smaller cocci and fewer tetrads than A. viridans does, it could be confused with pediococci or densely packed streptococci or enterococci. The most important routine tests are detection of leucine arylamidase, β-glucuronidase, PYR, hydrolysis of hippurate, and antibiotic susceptibility patterns.7,11,18,26,30,31 Rapid PYR testing is useful for distinguishing between A. viridans or enterococci (both PYR positive) and A. urinae (PYR negative). A Gram stain should be carefully examined for the characteristic arrangement in clusters and tetrads to rule out lactobacillus and other streptococcus. Pediococci are PYR negative and have a Gram stain morphology similar to that of A. urinae; however, they differ in their resistance to vancomycin and in their positive bile esculin test result.18,26 Transmission Electron Microscopy. The transmission electron microscope (TEM) operates on the same basic principles as the light microscope but uses electrons as “light source.” Their much lower wavelength makes it possible to get a resolution a thousand times better than with a light microscope. Cells of both A. urinae and A. viridans had a mean diameter of 0.90–1.10 μm, and strains of both species

214

showed a surface layer.32 In contrast to A. viridans, about 30% of all A. urinae cells showed signs of asymmetrical cell division.32 Sample preparation for TEM requires cultures in log phase. The cells can be fixed for 2 h in 3% glutaraldehyde in 0.1 M cacodylate buffer, pH 7.2, and then pelleted (8800 × g for 2 min). The precipitates should be enrobed in melted Noble agar (Difco) and small cubes with visible clusters of cells can be transferred to 3% glutaraldehyde and fixed overnight at 4°C. The specimens may be post fixed for 90 min in 1% (w/v) OsO4 in cacodylate buffer and stained with 2% (w/v) uranyl acetate in barbiturate buffer. The procedures for the dehydration, embedding in Epon (glycide ether) and the further preparation of thin sections can be performed as described by Blom et al.33 The stained and loaded samples may be viewed using TEM. Antimicrobial Susceptibility Tests. For this, the antimicrobial contained in a reservoir is allowed to diffuse out into the medium (e.g., blood agar) and interact in a plate freshly seeded with the test organisms. A variety of antimicrobial containing reservoirs are used, but the antimicrobial impregnated absorbent paper disc is by far the commonest type used (disc diffusion method). The results of in vitro antibiotic susceptibility testing guide clinicians in the appropriate selection of initial drugs used for individual patients in specific situations. The selection of an antibiotic panel for susceptibility testing is based on the commonly observed susceptibility patterns and is revised periodically. According to the antimicrobial susceptibility data of 30 aerococcal isolates obtained from Centers for Disease Control and Prevention,14 the MICs for nine strains were 0.5 g or more of penicillin per mL, with MICs for five strains being more than 1 g/mL; therefore, approximately 46% of aerococci tested were either relatively resistant or resistant to penicillin. Moreover, Christensen et al.19,28 documented that penicillin resistance should be the peculiar characteristic of A. viridans, capable of differentiating it from Aerococcus-like organisms. Table 20.1 shows the antibiotic susceptibilities of A. urinae, A. viridans, and A. sanguinicola. Fatty Acid Methyl Ester Analysis. Microorganisms can be classified based on gas chromatographic analysis of extracted microbial fatty acid methyl esters (FAMEs). Microbial fatty acid profiles are unique from one species to another, and this has allowed for the creation of very large microbial libraries. Location of the double-bond position of monounsaturated fatty acids of A. viridans was accomplished by combined gas chromatography (GC)–mass spectrometry analysis of dimethyl disulfide (DMDS) derivatives. The monoenoic fatty acids from whole bacterial cells were converted to methyl esters and then to DMDS adducts and analyzed by capillary GC–mass spectrometry.37,38 Multilocus Enzyme Electrophoresis (MLEE). Isolates are characterized by the relative electrophoretic mobilities of a large number of water-soluble cellular enzymes. Because the net electrostatic charge and, hence, the rate of migration of a protein during electrophoresis are determined by its amino acid sequence, mobility variants (electromorphs or

Molecular Detection of Human Bacterial Pathogens

TABLE 20.1 Antibiotic Susceptibilities of A. urinae and A. viridans. Data Has Been Adapted from Observations Reported Earliera Antibiotic Penicillin G Trimethoprimsulfamethoxazole Vancomycin Gentamicin Tetracycline Ciprofloxacin Ampicillin Erythromycin Cefuroxime Chloramphenicol

A. urinae

A. viridans

A. sanguinicola

S R

R S

S NA

S R S S S R S S

S S/R S R R R R R

S NA NA NA S S S NA

Abbreviations:  S, susceptible; R, resistant; NA, not available. Source: Uh, Y. et al., J. Korean Med. Sci., 17, 113, 2002; Zhang, Q. et al., J.  Clin. Microbiol., 38, 1703, 2000; Skov, R.L. et al., Diagn. Microbiol. Infect. Dis., 21, 219, 1995; Ibler, K. et al., Scand. J. Infect. Dis., 40, 761, 2008; Colakoglu, S. et al., Infection, 36, 288, 2008.

allozymes) of an enzyme can be directly equated with alleles at the corresponding structural gene locus.39 Christensen et al.32 used various enzymes for generating MLEE patterns and finally constructed UPGMA and Minimal Evolution trees based on the results. They found that the tree concurs with the ribotyping data and distinguished A. urinae strains as a group of strains distinct from the type strain of A. viridans. Sample preparation methods for MLEE have been adapted from those described by Selander et al.39 For this technique, whether cells are grown in broth or on agar plates is unimportant, since growth conditions do not affect the electrophoretic mobilities of enzymes. However, some enzymes may vary in activity (but not in mobility) on gels, depending on the type of medium in which the cells were grown. The activity level of certain enzymes may also vary with the composition of the culture medium, but, again, electrophoretic mobilities are not affected. Cells are normally lysed by sonication; however, any method of lysis that does not denature proteins may be used. After lysis and centrifugation at 30,000 × g for 20 min, aliquots of the several milliliters of lysate (supernatant) are transferred to three or four culture tubes and stored at −70°C until used for electrophoresis. At that temperature, lysates can be stored for several years without significant loss of activity of most enzymes. The stability of the enzymes varies markedly among species of bacteria, and so this step may be avoided. Apparatus for horizontal starch-gel electrophoresis is used. Starch gels are preferred over polyacrylamide gels. To prepare a gel, a suspension of starch and gel buffer is heated just beyond the boiling point.39 The suspension is immediately poured into a lucite gel mold. After the gel has cooled at room temperature for 2 h, it is wrapped in plastic film to prevent desiccation. Gels are used within 24 h of preparation. In loading a gel, pieces of Whatman no. 3 filter

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paper (9 mm by 6 mm) are individually dipped into samples of lysate, blotted on filter paper to remove excess liquid, and then inserted at 3-mm intervals in a continuous slit cut in the gel. Up to 20 lysates can be electrophoresed on a single gel. Pieces of filter paper dipped in amaranth dye are inserted at one or both ends of the slit to mark the migration front of the buffer line. During electrophoresis, a constant voltage is maintained, and the gel is cooled by a pan of ice supported above the gel mold on a thin plate of glass. Gels may be run in cold rooms or refrigerators at 4°C. Following electrophoresis, three or four horizontal slices (1–2 mm thick) are cut from the gel with a thin wire and incubated individually at 37°C in various selective enzyme staining solutions.39 The activity level and degree of separation of electromorphs of an enzyme may depend on the type and pH of the buffer system used and, to some extent, on the concentration of starch in the gel. Thus, an enzyme may appear to be monomorphic or polymorphic for only two or three electromorphs in one buffer system but exhibit 5–10 electromorphs in another buffer system. Occasionally, the relative mobilities of certain electromorphs are reversed in different buffer systems. Comparisons of the mobilities of enzymes from different isolates are made visually against one another on the same gel slice. It is not sufficient to compare relative mobilities of an enzyme in different strains by measuring distances of migration from the origin, even when the enzymes have been electrophoresed on the same gel. Each isolate is characterized by its combination of electromorphs over the number of enzymes assayed, and distinctive profiles of electromorphs, corresponding to unique multilocus genotypes, are designated electrophoretic types (ETs), which are equivalent to allele profiles. For analysis, a distance matrix in MEGA format may be generated using the computer program ETMEGA of T.S. Whittam (www.foodsafe.msu.edu/whittam/#programs) as mentioned by Chritensen et al.32 ETMEGA calculates the genetic distance between pairs of “electrophoretic types,” which were defined by distinct patterns of electrophoretic mobilities of the 11 housekeeping enzymes. Distance is measured as the proportion of mismatched loci between pairs of ETs. Null alleles are not used in the calculation of pair wise distances. Phylogenetic and molecular evolutionary analyses using this distance matrix may be conducted using MEGA version 4.32,40–42 Ribotyping. Ribotyping is a method that can identify and classify bacteria based upon differences in rRNA. It generates a highly reproducible and precise fingerprint that can be used to classify bacteria from the genus through and beyond the species level. DNA is extracted from a colony of bacteria and then restricted into discrete-sized fragments. In experiments performed by Christensen et al.32 ribotyping of the 26 A. urinae strains and the type strain of A. viridans with EcoR1 and HaeIII resulted in large DNA fragments of more than 40 kb, which could not be resolved using the RiboPrinter system. Ribotyping these strains with XbaI resulted in seven distinct ribotype patterns, each comprising between 1 and 12 strains. They showed that strains of A. urinae were clearly distinct from A. viridansT.

20.2  METHODS 20.2.1  Sample Preparation For extraction of Aerococcus DNA, a Wizard SV genomic DNA purification system (Promega, Madison, Wis.) may be used with some modifications. The bacterial pellet is resuspended in 400 μl enzymatic lysis solution (47 mM EDTA, 25 mg/ml lysozyme [Sigma, St. Louis, Mo.], 20 μg/ml lysostaphin [Sigma]) and incubated at 37°C for 2 h. Then, proteinase K (19.2 mg/ml; Roche) is added to a final concentration of 0.4 mg/ml and the mixture is incubated at 55°C for 1 h. Nuclei Lysis solution (Promega) and RNase solution (Promega) are added, and after mixing, the reaction solution is incubated at 80°C for 10 min. Further purification steps are done according to the instructions of the manufacturer. The final elution volume is 200 μl. For each PCR, 2 μl of this DNA extract is applied.

20.2.2  Detection Procedures (i) Aerococcus Viridans-Specific PCR  Martín et al. utilized A. viridans-specific primers AC2 (5′-GTGC TTGCACTTCTGACGTTAGC-3′) and AC4 (5′-TGAGC CGTGGGCTTTCACAT-3′) to amplify a 540 bp fragment from 16S rRNA gene.43 Furthermore, Grant et al. developed a rapid protocol for identification of A. viridans through PCR amplification of a 16S rRNA gene fragment, followed by detection with an oligonucleotide probe.44 (ii) PCR and Sequencing  Sequencing analysis of 16S rRNA gene allows identification of all Aerococcus species.45 In this protocol, a large fragment of the 16S rRNA gene (corresponding to positions 30 ± 1521 of the Escherichia coli 16S rRNA gene) is amplified by PCR using conserved primers close to the 3′ and 5′ ends of the gene. The PCR product is directly sequenced using a Taq Dye-Deoxy Terminator Cycle Sequencing kit (Applied Biosystems) and an automatic DNA sequencer. (iii) PCR Ribotyoing  PCR ribotyping was developed by Kostman et al.46 and Gurtler et al.47 in the 1990s as a response in part to the need in the clinical microbiology laboratory setting for expeditious epidemiological discrimination among pathogenic microorganisms without the use of probes, thus making the analysis more widely applicable.48 Primers complementary to the 3′ end of the 16S rRNA gene and the 5′ end of the 23S rRNA gene were used such that the amplified fragment revealed length heterogeneity of the internal spacer regions (ISR) located between the 16S and 23S genes of the six rRNA gene operons.49 However, PCR ribotyping proved not to be universally applicable.50 In some species, ISR length is variable within and between isolates, in others ISR lengths are limited to one or two sizes, usually dependent on the number of tRNAs present.48 In an attempt to further subtype isolates for which PCR ribotyping was nondiscriminatory, Ryley et al.51 and Shreve et al.52 performed PCR amplification of ISRs, as described above, followed by restriction endonuclease

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Molecular Detection of Human Bacterial Pathogens

Tissue or Cells

Quantitation by Nanodrop

Genomic DNA Extraction Genomic DNA Quantitation by Electrophoresis

Restriction digestion and size separation of DNA fragments

Hybridization with RNA probe

Dendrogram showing similarities between ribotypes

FIGURE 20.1  Demonstration of ribotyping. DNA is isolated from a cultured isolate, restricted and size separated in an agarose gel. The gel is then hybridized with labeled rRNA probe from a type strain of the species or genus, which binds to fragments containing copies of the rRNA operon. Following probe detection, fragments with bound probe are visualized, forming a characteristic RFLP. The advantages of ribotyping over other RFLP methods is that both highly variable and conserved sequences are highlighted by the probe, which further makes the method suitable to type strains over a broad phylogenetic range, and the same process can be used for analysis of strains within any species.

digestion. Christensen et al.32 found the following enzymes suitable for Aerococcus: XbaI, EcoR1, and HaeIII. An alternate variation of PCR ribotyping, amplified rRNA gene restriction analysis (ARDRA) is based on PCR amplification of the 16S rRNA gene followed by restriction digestion. Jayarao et al.53 developed this technique to determine subspecies of Streptococcus uberis, as a means of avoiding methods involving DNA hybridization or sequencing. Later, as described by Dupont Qualicon, the RiboPrinter automated ribotyping system came as a technological breakthrough with respect to convenience, reproducibility, and speed. The speed in this case appears to be two- to threefold greater than that for manual ribotyping, with far less labor-intensive procedures. For polyphasic characterization of A. urinae, automated ribotyping method (Figure  20.1) was used successfully with the Riboprinter (Qualicon Inc., Wilmington, DE).32

two distinct lineages of A. urinae exist, which can be identified by phenotypic markers. Future recognition of these biovars may reveal potential differences in ecology and clinical significance. In daily routine practice, species identification of Aerococcus causes problems. Routine diagnostic procedures should be able to identify this bacteria, and clinicians should be aware of the pathogenic potential of this organism.

ACKNOWLEDGMENTS We thank Dr. Mark Lawrence and Dr. Shane Burgess for their support in writing the manuscript. We are also grateful to Dr. Amit Ghosh, and Dr. Walter J. Diehl for encouragement.

REFERENCES

20.3  CONCLUSION The genus Aerococcus shows distinct interspecies differences, but constitutes a well-circumscribed genus as revealed by 16S rRNA sequence and other data. Phenotypic and molecular analyses demonstrate that A. urinae is to some extent a heterogeneous species belonging to the genus Aerococcus. At least



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Aerococcus







5. Lawson, P.A., Falsen, E., Ohlen, M., and Collins, M.D., Aerococcus urinaehominis sp. nov., isolated from human urine, Int. J. Syst. Evol. Microbiol., 51, 683, 2001. 6. Aguirre, M., and Collins, M.D., Phylogenetic analysis of some Aerococcus-like organisms from urinary tract infections: Description of Aerococcus urinae sp. nov., J. Gen. Microbiol., 138, 401, 1992. 7. Christensen, J.J. et al., Bacteremia/septicemia due to Aerococcus-like organisms: Report of seventeen cases. Danish ALO Study Group, Clin. Infect. Dis., 21, 943, 1995. 8. Christensen, J.J., Korner, B., and Kjaergaard, H., Aerococcuslike organism—an unnoticed urinary tract pathogen, Apmis, 97, 539, 1989. 9. Sierra-Hoffman, M., Watkins, K., Jinadatha, C., Fader, R., and Carpenter, J.L., Clinical significance of Aerococcus urinae: A retrospective review, Diagn. Microbiol. Infect. Dis., 53, 289, 2005. 10. Lawson, P.A., Falsen, E., Truberg-Jensen, K., and Collins, M.D., Aerococcus sanguicola sp. nov., isolated from a human clinical source, 51, 475, 2001. 11. Collins, M.D., Jovita, M.R., Hutson, R.A., Ohlen, M., and Falsen, E., Aerococcus christensenii sp. nov., from the human vagina, Int. J. Syst. Bacteriol., 49 Pt 3, 1125, 1999. 12. Collins, M.D., Jovita, M.R., Hutson, R.A., Ohlen, M., and Falsen, E., Note: Aerococcus christensenii sp. nov., from the human vagina, Int. J. Syst. Bacteriol., 49, 1125, 1999. 13. Christensen, J.J. et al., Aerococcus urinae: Intraspecies genetic and phenotypic relatedness, Int. J. Syst. Bacteriol., 47, 28, 1997. 14. Bosley, G.S. et al., Phenotypic characterization, cellular fatty acid composition, and DNA relatedness of aerococci and comparison to related genera, J. Clin. Microbiol., 28, 416, 1990. 15. Schuur, P.M.H. et al., Urinary tract infections with Aerococcus urinae in the south of The Netherlands, J. Urol., 161, 1414, 1999. 16. Martin, V. et al., Characterization of Aerococcus viridans isolates from swine clinical specimens, J. Clin. Microbiol., 45, 3053, 2007. 17. Razeq, J.H., Thomas, G.M., and Alexander, D., The first reported case of Aerococcus bacteremia in a patient with HIV infection, Emerg. Infect. Dis., 5, 838, 1999. 18. Facklam, R., and Elliott, J.A., Identification, classification, and clinical relevance of catalase-negative, gram-positive cocci, excluding the streptococci and enterococci, Clin. Microbiol. Rev., 8, 479, 1995. 19. Uh, Y., Son, J.S., Jang, I.H., Yoon, K.J., and Hong, S.K., Penicillin-resistant Aerococcus viridans bacteremia associated with granulocytopenia, J. Korean Med. Sci., 17, 113, 2002. 20. Kern, W., and Vanek, E., Aerococcus bacteremia associated with granulocytopenia, Eur. J. Clin. Microbiol., 6, 670, 1987. 21. Swanson, H., Cutts, E., and Lepow, M., Penicillin-resistant Aerococcus viridans bacteremia in a child receiving prophylaxis for sickle-cell disease, Clin. Infect. Dis., 22, 387, 1996. 22. Hermansson, K. et al., Structural studies of the capsular polysaccharide from Aerococcus viridans var. homari, Carbohydr. Res., 208, 145, 1990. 23. Nasoodi, A., Ali, A.G., Gray, W.J., and Hedderwick, S.A., Spondylodiscitis due to Aerococcus viridans, J. Med. Microbiol., 57, 532, 2008. 24. Tekin, A., Tekin, G., Turunc, T., Demiroglu, Z., and Kizilkilic, O., Infective endocarditis and spondylodiscitis in a patient due to Aerococcus urinae: first report, Int. J. Cardiol., 115, 402, 2007. 25. Ebnother, C., Altwegg, M., Gottschalk, J., Seebach, J.D., and Kronenberg, A., Aerococcus urinae endocarditis: Case report and review of the literature, Infection, 30, 310, 2002.

217 26. Zhang, Q., Kwoh, C., Attorri, S., and Clarridge, J.E., Aerococcus urinae in urinary tract infections, J. Clin. Microbiol., 38, 1703, 2000. 27. Augustine, T., Thirunavukkarasu, Bhat, B.V., and Bhatia, B.D., Aerococcus viridans endocarditis. Case report, Indian Pediatr., 31, 599, 1994. 28. Christensen, J.J., Korner, B., Casals, J.B., and Pringler, N., Aerococcus-like organisms: Use of antibiograms for diagnostic and taxonomic purposes, J. Antimicrob. Chemother., 38, 253, 1996. 29. Vandamme, P. et al., Polyphasic taxonomy, a consensus approach to bacterial systematics, Microbiol. Rev., 60, 407, 1996. 30. Christensen, J.J., Vibits, H., Ursing, J., and Korner, B., Aerococcus-like organism, a newly recognized potential urinary tract pathogen, J. Clin. Microbiol., 29, 1049, 1991. 31. Kristensen, B., and Nielsen, G., Endocarditis caused by Aerococcus urinae, a newly recognized pathogen, Eur. J. Clin. Microbiol. Infect. Dis., 14, 49, 1995. 32. Christensen, J.J. et al., Aerococcus urinae: Polyphasic characterization of the species, Apmis, 113, 517, 2005. 33. Blom, J., Mansa, B., and Wilk, A., A study of Russell bodies in human monoclonal plasma cells by means of immunofluorescence and electron microscopy, Acta Pathol. Microbiol. Scand., A 84, 335, 1976. 34. Skov, R.L., Klarlund, M., and Thorsen, S., Fatal endocarditis due to Aerococcus urinae, Diagn. Microbiol. Infect. Dis., 21, 219, 1995. 35. Ibler, K. et al., Six cases of Aerococcus sanguinicola infection: Clinical relevance and bacterial identification, Scand. J. Infect. Dis., 40, 761, 2008. 36. Colakoglu, S. et al., Three cases of serious infection caused by Aerococcus urinae: A patient with spontaneous bacterial peritonitis and two patients with bacteremia, Infection, 36, 288, 2008. 37. Moss, C.W., and Daneshvar, M.I., Identification of some uncommon monounsaturated fatty acids of bacteria, J. Clin. Microbiol., 30, 2511, 1992. 38. Moss, C.W., Wallance, P.L., and Bosley, G.S., Identification of monounsaturated fatty acids of Aerococcus viridans with dimethyl disulfide derivatives and combined gas chromatography-mass spectrometry, J. Clin. Microbiol., 27, 2130, 1989. 39. Selander, R.K. et al., Methods of multilocus enzyme electrophoresis for bacterial population genetics and systematics, Appl. Environ. Microbiol., 51, 873, 1986. 40. Kumar, S., Nei, M., Dudley, J., and Tamura, K., MEGA: A biologist-centric software for evolutionary analysis of DNA and protein sequences, Brief Bioinform., 9, 299, 2008. 41. Kumar, S., Tamura, K., and Nei, M., MEGA3: Integrated software for Molecular Evolutionary Genetics Analysis and sequence alignment, Brief Bioinform., 5, 150, 2004. 42. Tamura, K., Dudley, J., Nei, M., and Kumar, S., MEGA4: Molecular Evolutionary Genetics Analysis (MEGA) software version 4.0, Mol. Biol. Evol., 24, 1596, 2007. 43. Martín, V. et al., Characterization of Aerococcus viridans isolates from swine clinical specimens, J. Clin. Microbiol., 45, 3053, 2007. 44. Grant, K.A. et al., Rapid identification of Aerococcus viridans using the polymerase chain reaction and an oligonucleotide probe, FEMS Microbiol. Lett., 95, 63, 2006. 45. Lawson, P.A. et al., Aerococcus urinaehominis sp. nov., isolated from human urine, Int. J. Syst. Evol. Microbiol., 51, 683, 2001.

218 46. Kostman, J.R. et al., A universal approach to bacterial molecular epidemiology by polymerase chain reaction ribotyping, J. Infect. Dis., 171, 204, 1995. 47. Gurtler, V., and Stanisich, V.A., New approaches to typing and identification of bacteria using the 16S-23S rDNA spacer region, Microbiology, 142, 3, 1996. 48. Bouchet, V., Huot, H., and Goldstein, R., Molecular genetic basis of ribotyping, Clin. Microbiol. Rev., 21, 262, 2008. 49. Gurtler, V., and Barrie, H.D., Typing of Staphylococcus aureus strains by PCR-amplification of variable-length 16S-23S rDNA spacer regions: characterization of spacer sequences, Microbiology, 141, 1255, 1995. 50. Cartwright, C.P., Polymerase chain reaction ribotyping: A “universal” approach? J. Infect. Dis., 172, 1638, 1995.

Molecular Detection of Human Bacterial Pathogens 51. Ryley, H.C., Millar-Jones, L., Paull, A., and Weeks, J., Characterisation of Burkholderia cepacia from cystic fibrosis patients living in Wales by PCR ribotyping, J. Med. Microbiol., 43, 436, 1995. 52. Shreve, M.R., Johnson, S.J., Milla, C.E., Wielinski, C.L., and Regelmann, W.E., PCR ribotyping and endonuclease subtyping in the epidemiology of Burkholderia cepacia infection, Am. J. Respir. Crit. Care. Med., 155, 984, 1997. 53. Jayarao, B.M., Dore, J.J., Jr.. and Oliver, S.P., Restriction fragment length polymorphism analysis of 16S ribosomal DNA of Streptococcus and Enterococcus species of bovine origin, J. Clin. Microbiol., 30, 2235, 1992.

21 Bacillus Noura Raddadi, Ameur Cherif, and Daniele Daffonchio CONTENTS 21.1 Introduction.......................................................................................................................................................................219 21.1.1 Classification, Morphology, and Biology of Bacillus spp. ...................................................................................219 21.1.2 Genetic Baseline of B. cereus Group Ecotypes.................................................................................................... 220 21.1.3 Species Discrimination in B. cereus Group......................................................................................................... 220 21.1.4 Clinical Features and Toxicology of Human Pathogenic Bacillus spp................................................................ 220 21.1.4.1 Bacillus anthracis.................................................................................................................................. 220 21.1.4.2 Bacillus cereus and Other Human Pathogenic Bacillus spp. ............................................................... 221 21.1.5 Diagnosis Techniques........................................................................................................................................... 223 21.1.5.1 Conventional Diagnosis Techniques...................................................................................................... 223 21.1.5.2 Molecular Diagnosis Techniques........................................................................................................... 223 21.2 Methods............................................................................................................................................................................ 224 21.2.1 Sample Preparation............................................................................................................................................... 224 21.2.2 Detection Procedures............................................................................................................................................ 224 21.3 Conclusions and Future Perspectives............................................................................................................................... 225 References.................................................................................................................................................................................. 226

21.1  INTRODUCTION 21.1.1  C  lassification, Morphology, and Biology of Bacillus spp. Bacteria in the genus Bacillus are gram-positive, spore formers, and rod-shaped. They are very diverse in terms of physiology, ecological niche, genes sequences, and regulation. Their impact on human activity varies from probiotic effect1,2 to severe pathogenecity. The oldest, most infectious, and potentially lethal human disease is the anthrax that is caused by Bacillus anthracis.3 However, many other species within this genus have emerged as new human pathogens associated with foodborne diseases that can cause severe and even fatal infections. These include B. cereus, B. weihenstephanenesis, B. pumilus, B. mojavensis, B. licheniformis, B. subtilis, and B. circulans. All Bacillus species, due to endospore formation, can survive heat treatment and disinfection procedures and hence pose serious health concerns as contaminants of food or in hospital settings as a cause of nosocomial infections,4–10 as well as severe illnesses, especially in immunocompromised patients or immunocompetent individuals with risk factors such as intravenous drug use, hemodialysis, and leukemia. The production of toxins and enzymes by these bacteria is considered the causative agent of several human diseases. In this chapter, we survey the different human pathologies caused by Bacillus spp., mainly those of the B. cereus group.

Before giving details on the clinical features, toxicology, and the methods of detection of these bacteria and their corresponding pathogenecity factors, we will briefly introduce the complex ecology and the identification challenges that characterize several Bacillus species. The B. cereus group, which shows a “bivalent face” in terms of its important impact on human activity will be used as a paradigm.11,12 B. anthracis is the most serious human pathogen in this group and the major subject of this chapter. This bacterium is the etiological agent of anthrax, a potentially fatal disease for humans and animals, and a bioterrorism agent. B. cereus is the most known foodborne pathogen that has been associated with food-poisoning illnesses13,14 and other kinds of clinical infections that are reported in this chapter. B. thuringiensis is an insect pathogen widely used as biopesticide, and differently from B. cereus, has a very useful impact on human activities being widely used in agriculture for insect/pest biocontrol. B. weihenstephanensis, a psychrotolerant species capable of growing at temperatures as low as 4°C–6°C is implicated in food spoilage.15 In addition to B. anthracis, B. cereus, B. thuringiensis, and B. weihenstaephanenesis, the B. cereus group encompasses two other species, B. mycoides and B. pseudomycoides, which are typically isolated from soil and plant rhizosphere. The six species are known to be strictly related phylogenetically, as has been shown by DNA–DNA hybridization studies16–18 and the sequencing of the ribosomal RNA 219

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genes.19–21 However, a marked variability is always observed when large collections of strains are examined by DNA fingerprinting methods that target the whole genome22–26 and/or discrete genes.27–31 Hence, the phylogenetic and taxonomic relationships among these species are still open to debate. The genetic variability observed within and between these species has raised several interesting questions and poses a challenge to microbiologists: (i) What is the evolutionary pathway of these species and how are they differentiated during evolution? (ii) What is the genetic baseline driving the different ecotypes and the virulence?

21.1.2  Genetic Baseline of B. cereus Group Ecotypes The ecology of these bacteria is still far from clear, despite several recent analyses that have shown that a major environmental niche for these bacteria is the invertebrate intestinal tract.32–34 In a general model, these species commonly live associated to the intestine of invertebrates, assuming a symbiotic life style. But, occasionally, they escape such an ecotype, becoming invasive for other animal hosts that are in specific insects, arthropods, and nematodes for B. thuringiensis, and in mammals and humans for B. anthracis and B. cereus.35 Apart from this general scheme, the ecology of these species outside the invertebrate host is not completely clear, and the ability of several environments with which these bacteria have been commonly associated, to continuously support their life cycle has been questioned. It has been shown that with respect to other soil-inhabiting bacteria, such as those of the B. subtilis group that have genomes harboring many genetic determinants for the metabolism of plant-derived sugars, B. cereus group bacteria are rich in genes for protein metabolism. This suggests that they evolved in relation to nutrient-rich environments, such as animal guts or animal tissues and fluids, rather than the plant environment.36,37 Specialization on different “animal” niches has lead to different evolution of the species in the B. cereus group. For instance, B. anthracis has been proposed to diverge from the other members of the group by specializing as a lethal mammal pathogen. It has been supposed that B.  anthracis has evolved along two possible pathways38: In the first pathway, one considers B. anthracis to be a relatively ancient organism with a low growth rate, determined by the previously mentioned ecological constraints, and evolving separately from a common ancestor with B. cereus and other relatives. In the second pathway, it has been supposed to be derived relatively recently from B. cereus through the acquisition and rearrangement of plasmids resulting in the actual pXO plasmid pattern responsible for lethal virulence.

21.1.3  Species Discrimination in B. cereus Group The similarity among such closely related bacteria as the species of the B. cereus group posed serious challenges for species discrimination. This is an important point to address, considering the differences in the virulence potential of these microorganisms. Typically, B. anthracis can be

Molecular Detection of Human Bacterial Pathogens

differentiated from most of B. cereus and B. thuringiensis strains by specific tests, such as penicillin G and gamma phage sensibility, lack of motility, and beta-hemolysis on blood agar. However, a problem still exists for isolates borderline between species, such as, for instance, the pathogenic B. cereus G9241, for which these tests fail to recognize its pathogenic potential.39 This strain has been confirmed to be a B. cereus, harboring a virulence plasmid very similar to plasmid pXO1 of B. anthracis and a second plasmid encoding for a capsule synthesis. However, such a second plasmid and the capsule-coding genes were completely different from the pXO2 plasmid and the typical capsule of B. anthracis.39 Thus, considering the virulence potential of strains that are genetically the near neighbors of B. anthracis, approaches that might rapidly identify these strains are of great interest. From a safety perspective, these strains could represent alternative hosts for B. anthracis toxin genes.39 Several B. cereus strains resulted, strictly related to B. anthracis. For example, Keim et al.38 and Radnedge et al.40 individuated by amplified fragment-length polymorphism (AFLP) B. cereus and B. thuringiensis strains closely related to B. anthracis. Radnedge et al.41 tried to identify, by suppression subtractive hybridization, genomic regions of B. anthracis absent in these closely related strains. Apart from AFLP, several other methods based on whole-genome fingerprinting have been used for typing B. anthracis and for the identification of species borderline strains, like, among others, rep-PCR12 and multilocus sequence typing (MLST)42 or, recently, comparative genomics43 and microarray analysis.44 Alternative approaches have been based on length or sequence polymorphisms in variable‑number tandem repeats in multiple loci (multilocus VNTR analysis [MLVA])45 or on signature single nucleotide polymorphisms (SNPs) in the genome.46 MLVA has been used as a gold standard for subtyping B. anthracis isolated worldwide (see among others35,45,47,48). Together with MLVA, SNP analysis is greatly contributing in typing and tracing B. anthracis isolates, especially by the whole-genome SNP analysis.49 Besides those identified in the whole genome, SNPs in housekeeping genes have been nicely exploited to identify strains related to certain species or genetic types. For example, Prüss et al.50 showed that certain nucleotides in the 16S rRNA gene and their relative prevalence among the different ribosomal operons in the genome correlate with the psychrotolerance of B. cereus strains, and hence, are signature of the species B. weihenstephanensis. Gierczyn´ski et al.51 developed a simple and cost-effective restriction-siteinsertion PCR (RSI-PCR)–based assay that precisely detects the B. anthracis-specific nonsense mutation in plcR gene by restriction digestion with the endonuclease SspI.

21.1.4  C  linical Features and Toxicology of Human Pathogenic Bacillus spp. 21.1.4.1  Bacillus anthracis B. anthracis, the causative agent of anthrax disease, has been well studied for over 150 years and continues to be a threat as a biological weapon.3 This bacterium, forms spores (1–2

Bacillus

μm in size) able to withstand several kinds of stresses that are usually lethal to vegetative cells, such as ultraviolet and ionizing radiation, heat, various chemicals,52 and oxidative stress.53 Anthrax is primarily a disease affecting herbivores, but it can be transmitted to all mammals, including humans. Typically, animals become infected with B. anthracis by ingesting spores while grazing on contaminated soil or feed and also through the skin via biting insects.54 Transmission of the spores to humans occurs through three routes of entry: (i) cutaneously, by direct contact with infected animals or by handling infected animal products55; (ii) respiratory, by inhalation of sufficient quantity of spores; and (iii) oropharyngeal and/or gastrointestinal (GI), by the ingestion of undercooked, contaminated meat.56 All three forms of anthrax can be fatal if not treated in time, but the cutaneous form is often self-limiting.57 A typical anthrax skin lesion corresponds to an ulcer covered by a characteristic black eschar.58 The pathogenecity of B. anthracis depends on two major factors: a poly-γ-d-glutamic acid capsule and the three-component secreted anthrax toxin codified by plasmids pXO2 and pXO1, respectively. In addition to protecting the bacteria from the phagocytosis, the capsule allows bacterial proliferation and subsequent toxemia to develop in the bloodstream because of its poor immunogenic properties. The anthrax toxin is composed of three proteins known as protective antigen (PA), lethal factor (LF), and edema factor (EF). Once in the body, the PA binds to receptors that are present on most cells, including macrophages, and after proteolytic activation, it combines to LF and EF, allowing their delivery inside the cell.59 Transcription of the genes pag, lef, and cya encoding the three-toxin proteins PA, LF, and EF, respectively, is regulated by a gene known as the anthrax toxin activator (atxA); whereas the capsule proteins, codified by the capBCADE operon, are regulated by AcpA and AcpB.60,61 Once in the host and after their phagocytosis by local macrophages, spores germinate and multiply in their way to the regional lymph nodes.41 Along with other mechanisms, two spore-bound superoxide dismutases (SOD15 and SODA1), enzymes that are capable of detoxifying oxygen radicals,62 were recently reported to play a crucial role in protecting the spores against the oxidative burst of macrophages in mouse model.53 Following spore germination and bacterial proliferation, the capacity of the lymph nodes is exceeded; hence, the vegetative bacteria enter the bloodstream, which leads to bacteremia and the distinctive pathology of edema, hemorrhage, and tissue necrosis.59 Dissemination of B. anthracis by hematogenous or lymphatogenous spread from a cutaneous, mediastinal, or GI focus can lead to infection at other sites, such as the brain, allowing entry into the central nervous system (CNS) and development of anthrax meningitis.63 Another form of anthrax meningitis in which no primary focus is found has been reported and is known as “primary anthrax meningitis.”64 With regard to the epidemiology, anthrax infection has been reduced dramatically in developed countries as the result of the widespread use of vaccines in high-risk people

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and animals, as well as the improvements in industrial hygiene. Currently, only very few cases of human anthrax occur naturally in these countries. However, the scenario is different in developing countries such as Africa and Asia, where the disease is endemic; this is due in part to difficulties of implementing vaccination programs. Of the three forms of the disease, cutaneous anthrax is the most frequent, followed by the inhalational and GI or oropharyngeal form.55,57,65,66 Only few case reports have been published regarding anthrax meningitis epidemiology, and most of them report anthrax meningitis in settings of one of the three forms of the disease.57,64,67,68 21.1.4.2  Bacillus cereus and Other Human Pathogenic Bacillus spp. Due to their ubiquitous presence in the environment, Bacillus spp., other than B. anthracis, isolated from clinical specimens have been often considered as a contamination; hence, most laboratories do not identify these isolates further. It has been reported that Bacillus spp. are isolated from 0.1% to 0.9% of blood cultures, but only 5%–10% of these are clinically significant infections.69–71 However, in recent years there has been a growing number of reports dealing with their implication in severe and even lethal infections. Bacillus spp. other than B. anthracis were shown to cause nosocomial bacteremia, pneumonia, brain and liver abscesses, infection of CNS, periodontitis, peritonitis, endophthalmitis, soft-tissue infection, and meningitis. Nosocomial Bacteremia. B. cereus is the most frequently isolated of Bacillus spp. involved in the development of nosocomial infections.7–10,72,73 Van Der Zwet72 described an outbreak of invasive B. cereus infections in a neonatal intensive care unit caused by contaminated balloons used for manual ventilation. These balloons had been cleaned with detergent, which was insufficient to kill spores of B. cereus. Kalpoe et al.9 reported an outbreak of vancomycinresistant B. cereus respiratory tract colonization in pediatric intensive care unit (PICU) patients due to contamination of reusable ventilator air-flow sensors. B. cereus spore contamination of alcohol used to disinfect the air-flow sensors was postulated to be the source of the dissemination. Dohmae et al.8 reported on five bloodstream infections in five patients related to catheter use. B. cereus contamination of reused towels, which would have occurred during the wash process, was considered to be the origin of this outbreak. Ohsaki et al.7 reported on the occurrence of three consecutive cases of B. cereus bacteremia in patients with hematologic malignancies after hospital renovation. The authors considered that filters used in the heating, ventilation, and air-conditioning systems, as well as towels and gowns, were the probable sources of the B. cereus outbreak. Kuroki et al.,10 demonstrated that B. cereus and B.  thuringiensis species previously implicated in nosocomial bacteremia by catheter infection were able to form biofilm after 24 h of incubation. To date, there is one report describing the occurrence of fatal B. thuringiensis bacteremia, in a neutropenic patient who suffered from severe pulmonary disease,74 indicating

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that this entomopathogenic bacterium may behave as an opportunistic pathogen in immunocompromised individuals. B. cereus together with B. licheniformis and B. pumilus were shown to be implicated in 3.4% of 350 bacteremic episodes that occurred during a 5-year period in patients having hemotological malignancies.75 To date, only one case of B. licheniformis nosocomial infection, affecting a very-low-birth-weight neonate in a neonatal intensive care unit, has been reported.76 In immunocompetent individuals, B. licheniformis bacteremia has been reported in association with central venous catheters.77 B. pumilus has also been implicated in persistent bacteremia in a patient with cholangitis and in nosocomial bacteremia via catheter infection.78,79 In addition to antibiotic treatment, the central venous catheter may need to be removed for complete cure. Taken together, these reports indicate that, although cases of Bacillus spp. other than B. anthracis and B. cereus bacteremia are rare in healthy individuals, it is important to consider the potential pathogenicity of Bacillus spp. in immunocompromised hosts, including premature infants and neonates, as well as in patients with various risk factors. Endophthalmitis. Infectious endophthalmitis, the inflammation of intraocular tissues or fluids due to intraocular infection, is a major public health concern. In addition to fungi and gram-negative bacteria80; different Bacillus species, such as B. megaterium,81 B. subtilis, B. mycoides, B. pumilus, B.  flexus, and B. thuringiensis, have been reported as causative agents of infectious endophthalmitis.82 However, the most important eye pathogen in this genus is B. cereus because of its ability to blind rapidly. Endophthalmitis associated to this bacterium often results in significant vision loss or loss of globe architecture in 1–2 days.5,83 The eye pathogenesis of B. cereus is due to rapid growth, migration, and toxin production by this microorganism, which lead to the failure of the host immune defenses.82,84 In addition to toxin production by the offending bacterium, the pathogenesis of B. cereus was also attributed to the intraocular inflammatory response and, in particular, to tumor necrosis factoralpha (TNFα) in experimental endophthalmitis.85 Recently, B. cereus was also shown to induce the permeability of the blood-ocular barrier in experimental endophthalmitis.86 Foodborne Illnesses. Bacillus cereus is well known as a common cause of emetic and diarrheal food poisoning. The symptoms of these foodborne illnesses are usually mild and the disease is often self-limiting. However, this opportunistic human pathogen can be associated with severe intoxications and infections that can be fatal, even in otherwise healthy individuals. Indeed, three fatal cases due to fulminant liver failure following intoxication with cereulide-producing B. cereus strains and liver abscess have been reported to date.13,87,88 Other Bacillus spp. have also been implicated in foodborne illnesses, including B. weihenstephanensis, B. subtilis, B. licheniformis, and B. pumilus; while other species, such as B. mojavensis, B. firmus, B. megaterium, B. simplex, and B. fusiformis were shown to produce toxins with a cereulide-like mode of action.14

Molecular Detection of Human Bacterial Pathogens

Pneumonia. Bacillus cereus has also been associated with fulminant pneumonia in five otherwise healthy metalworkers: a single nonfatal case in 1994 and four fatal cases: two in 1997 and two in 2003.89–91 The B. cereus strains responsible for these cases of fulminant pneumonia were shown to produce a capsule and to have the genetic determinants that codify for the anthrax toxins.92 A sixth case of fatal pneumonia caused in a leukemic patient by carbapenem-resistant B. cereus has also been reported recently.93 The fact that B. cereus has been implicated in such unusual severe fatal pneumonia in immunocompetent individuals highlights the need for increased awareness of the potential for this opportunistic human pathogen to cause serious systemic infections, even in patients who appear to be otherwise healthy. Thus, a better understanding of the environmental and occupational risk factors that may increase the susceptibility of individuals to infection by this bacterium should be considered. Periodontitis. The genus Bacillus has received relatively little attention in oral microbiology, and the Bacillus spp. isolated from human oral samples have been often regarded as transient microflora or as external contaminants. However, there have been several reports on the implication of Bacillus species, such as B. cereus/B. thuringiensis,94 B. pumilus,95 biofilm-forming B. subtilis,96 B. circulans, B. laterosporus, B. lentus, and B. sphaericus97 in oral disease, mainly gingivitis and periodontitis. On the other hand, cryptic (having no definable origin) pyogenic infections of the central nervous system (CNS) secondary to dental affections have been reported in which B. subtilis and B. circulans are implicated.98 These cases indicate that, although dental and paradental infections seem to be a rare cause of intracerebral or intraspinal infections in otherwise healthy individuals, it is recommended that a dental focus should always be considered in the evaluation and treatment of these kinds of cryptic infections of the CNS. Meningoencephalitis and Meningitis. In addition to B. anthracis, to the best of our knowledge, only B. cereus has been reported to be implicated in cases of meningoencephalitis in preterm infants99 and neonates100 or in cases of meningitis in neonates101,102 and immunocompromised individuals.103,104 Soft-tissue Infection. To date, there have been two case reports that have implicated B. cereus in necrotizing fasciitis and myositis that mimick clostridial myonecrosis in immunocompromised patients.105,106 This is a severe infection of skin and soft tissue by B. cereus that has symptoms similar to clostridial gas gangrene: in both cases the infection is caused by a gram-positive rod, and gas production and muscles inflammation (myositis) could be observed. However, to select the appropriate therapy, it is very important to differentiate between the two kinds of infections. Indeed, the treatment of clostridial gas gangrene is usually based on penicillin, an antibiotic to which B. cereus is resistant because it produces β -lactamase. B. cereus has to be suspected as a pathogen when the skin and soft-tissue infection occurs with sepsis in immunocompromised patients; whereas, Clostridium have to

Bacillus

be considered when the infection occurs after trauma in otherwise healthy individuals. Another criterion for the differentiation between the two conditions is the site of infection: for clostridial gas gangrene, the primary site of inflammation is the muscle parenchyma, in contrast to B. cereus infection, where inflammation often develops from the subcutaneous tissues to muscles.106 Peritonitis. B. cereus,107,108 B. circulans,109 and B. liche­ niformis110 have also been implicated in peritonitis in patients  undergoing peritoneal dialysis. There is also one case report on B. cereus peritonitis after cesarean section.111

21.1.5  Diagnosis Techniques The diagnosis of a human illness is based on the combination of different clues including the symptoms of the illness, the history of the patient, the possible risk factors, and the detection and identification of the suspected agent and/or its pathogenicity factors. Conventional detection procedures of Bacillus human pathogens are often based in the first step on the isolation and identification of the bacterium on selective cultural media from clinical specimens based on biochemical, phenotypic, and physiological characteristics. Otherwise, detection of the whole microorganism or its pathogenecity factors could be performed by immunological assays. Nucleic acid–based approaches are, however, most-often applied due to their sensitivity and rapidity. 21.1.5.1  Conventional Diagnosis Techniques Fast and reliable identification of the pathogenic microorganism implicated in the illness is key to a timely and adequate treatment of a patient. Typically, B. anthracis can be identified from clinical specimens and differentiated from other Bacillus spp. by classical microbiological tests that are recommended by the World Health Organization (WHO) and by the Centers for Disease Control and Prevention (CDC). Like the other species of the B. cereus group, B. anthracis is positive to Gram staining and produces spores; however, it can be distinguished from the nonanthrax group (B. cereus and B. thuringiensis) by its capacity to produce a polypeptide capsule in vivo and in vitro when grown under suitable conditions, its sensitivity to gamma phage and penicillin G, and by the lack of motility and β-hemolytic activity on sheep or horse blood agar medium.57 In addition to blood agar medium, B. anthracis can be isolated on the selective polymyxin-lysozyme-ethylenediaminetetraacetic acid-thallium acetate (PLET) agar112 or on anthracis chromogenic agar (ACA).113 On PLET medium, B.  anthracis colonies appear small, white, domed, and circular. Anthracis chromogenic agar contains the chromogenic substrate 5-bromo-4-chloro-3-indoxyl-choline phosphate that yields teal colonies upon hydrolysis by phosphatidylcholine-specific phospholipase C (PC-PLC), also known as lecithinase C. Upon incubation at 35°C–37°C, B. cereus and B. thuringiensis colonies are dark teal-blue within 24 h; whereas, B. anthracis colonies need at least 48 h to develop

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this color because of the slower rate of synthesis of this enzyme as a result of the truncated plcR gene and PlcR regulator. However, the blood-agar medium is easier and more cost effective, and hence is recommended for the isolation of B. anthracis from samples not overloaded with background bacilli, such as clinical specimens. The selective media PLET and ACA are suitable for the isolation of B. anthracis by well trained and expert personal, from samples with high numbers of bacteria but taking into account that there are limitations.114 However, this classical identification procedure is timeconsuming and needs at least 24–48 h to be accomplished. In addition, there are several other challenges in the detection of B. anthracis: (i) not all the B. anthracis isolates fit all the above described characteristics, and atypical B.  anthracis isolates lacking pXO2 (and hence capsule) have been identified; (ii) pathogenic capsule-producing B. cereus strains harboring plasmids similar to those of B. anthracis have been identified in cases of human fatal pneumonia and from the carcasses of great apes in Africa115; and (iii) blood cultures do provide B. anthracis identification, whereas cultures from skin or lesions are not reliable as they yield positive results in only 60%–65% of cases, especially after antibiotic treatment.58 The limitations of culture-based diagnosis of pathogenic Bacillus spp. causing human illness make necessary the application of quicker and more specific approaches, especially for the detection of B. anthracis. Several antibodybased tests targeting spores, vegetative cells, and anthrax toxin proteins have been developed for the specific detection of B. anthracis.59,116–118 Recently, several other approaches targeting the B. anthracis capsular antigen detection by latex agglutination119 or the entire organism mediating phage bioluminescence120 have been reported. However, these antibody-based assays bear the limitation of sensitivity since low concentration of antigens could not be detected in the first steps of bacterial infection. 21.1.5.2  Molecular Diagnosis Techniques DNA isolation from clinical specimens is a critical step in molecular biology–based diagnosis techniques because of the presence of high levels of inhibitors and host DNA in relation to the amount of DNA from the pathogenic bacterium, especially in the first step of the infection. Also, safety is an important consideration for the personnel handling specimens that are suspected to contain infectious B. anthracis spores, which are known to be difficult to inactivate. Several extraction kits are commercially available for the rapid isolation of DNA to be used in nucleic acid–based detection assays. In particular, the UltraClean Microbial DNA Isolation Kit (Mo Bio Laboratories, Inc.) was shown to offer the best method for depleting viable B. anthracis spores from samples.121 Dauphin et al.122 reported that gamma irradiation of DNA preparations allowed inactivation of residual B. anthracis spores; their complete removal could be reached by centrifugal filtration with 0.1-micron filter unit.123

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21.2  METHODS 21.2.1  Sample Preparation Bacterial Isolation. Bacillus spp. are often isolated from clinical specimens on 5%–7% horse or sheep blood agar medium. After overnight incubation at 35°C, nonanthrax bacteria of the B. cereus group are gram-positive, rodshaped, spore formers, motile, and beta-hemolytic; whereas B. anthracis is nonmotile and nonhemolytic. B. anthracis is also sensitive to gamma phage and penicillin G. Media like PLET or ACA could also be used for the selective isolation of B. anthracis. DNA Isolation. DNA template for PCR reactions can be prepared from the purified isolates by a simple boiling method as following. Cells are collected by centrifugation (5000 × g for 2 min) from 1 mL of a 4 h-culture grown in nutrient broth at 35°C. The cell pellet is washed once with 500 µL of sterile MilliQ water or TE buffer (pH = 8), centrifuged, resuspended in 100 µL of sterile MilliQ water or TE buffer, and boiled for 10–15 min. The boiled samples are centrifuged at 10,000 × g for 5 min to precipitate cell debris, and the supernatants, which contain nucleic acids, are collected and stored at −20°C until use. There are also several commercial kits available for total DNA extraction from overnight-grown pure bacterial cultures as well as from spores.121 The phenolchloroform extraction protocol is also useful to obtain pure high-molecular-weight genomic DNA.124 Total DNA from clinical specimens can be isolated using commercial DNA extraction kits, such as the ChargeSwitch gDNA Mini Bacteria Kit (Invitrogen); DNeasy Blood and Tissue kit (Qiagen); NucliSens Isolation Kit (Biomerieux); Puregene Genomic DNA Purification Kit (Qiagen); QIAamp DNA Blood Mini Kit (Qiagen); MagNA Pure LC (MPLC) microbiology kit MGrade (Roche Diagnostics GmbH); MagSi-DNA isolation kit for blood (Magna-Medics Diagnostics B.V.), and the UltraClean Microbial DNA Isolation Kit (Mo Bio Laboratories), following the manufacturers’ instructions. However, this step of sample processing may be hampered by the low concentration of the pathogen and the dominance of human DNA, especially in samples such as blood, cerebrospinal fluid, or in other primary sterile sites. Horz et al.125 developed two new kits, MolYsis® (Molzym GmbH & Co. KG, Bremen, Germany) and Pureprove® (SIRS-Lab GmbH, Jena, Germany) which is also known as Looxster® Universal kit, for the enrichment of bacterial DNA from blood samples that were also shown to be efficient using oral samples from plaque periodontitis and caries. Hansen et al.126 found that pretreatment of blood samples with MolYsis Basic prior to DNA isolation with MolYsis Complete Kit improves the detection of the DNA of pathogenic bacteria from blood samples allowing a detection limit of 50 CFU/mL in multiplex real-time PCR.

21.2.2  Detection Procedures Culture-dependent techniques for direct isolation and identification of Bacillus spp. based on traditional methods,

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looking to metabolism and physiology, is time consuming and laborious. Also, it is not possible to rely on bacterial culture for the diagnosis of cutaneous human anthrax, for which it has been reported that the bacterial culture– based diagnosis yields positive results only in 60%–65% of cases.58 Furthermore, the presence of atypical nonencapsulated B. anthracis strains and pathogenic capsule-producing B. cereus isolates represents a further challenge for a specific and accurate identification based on this phenotypic character. Thus, a combination of classical microbiological assays and nucleic acid–based methods is often necessary to confirm the identity of the infectious bacterial agent. Actually, there is a wide variety of molecular-based techniques for the detection of B. anthracis, including PCR, real-time PCR, and multiplex PCR. All these techniques are based on the specific amplification of the genes pag, lef, and cap. However, all these genetic determinants reside on plasmids; hence B. anthracis isolates that lack one of the plasmids can not be easily identified based on these techniques. For this reason, chromosomal-based B. anthracis–specific markers, such as BA813, rpoB, gyrA, gyrB, saspB, plcR, and BA5345 have been identified.51,127–129 Recently, Antwerpen et al.129 reported the development of a TaqMan real-time–based assay that targets the locus BA_5345, and that specifically detected B. anthracis, as was shown by testing 328 Bacillus strains. Within this sequence, they amplified a 96-bp large fragment by using the primers dhp61_183- 113F (5′-CGTAAGGACAATAAAAGCCGTTG T-3′), dhp61_183–208R (5′-CGATACAGACATTTATTGGG AACTACAC-3′) and TaqMan -probe dhp61_183–143T (5′-6FAM-TGCAATCGATGAGCTAATGAACAATGACCCT– TAMRA-3′). The TaqMan® PCR assay is performed in a 20  µL PCR-reaction mixture containing 1× ABITaqMan® mix (Applied Biosystems, Darmstadt, Germany); 0.9 µM primer dhp61_183–113F; 0.3 µM of primer dhp61_183–208R and 0.25 µM of labeled probe dhp61_183–143T; and 2 µL of template DNA. Amplification is carried out on the MX3000 Real-Time PCR System (Stratagene) with thermal conditions as follows: an initial denaturation step of 10 min at 95°C is followed by 45 cycles consisting of 15 s at 95°C and 1 min at 55°C. This assay is reported to have a detection limit of 12.7 copies/reaction. Another molecular assay that can be applied for the rapid and sensitive/quantitative detection of B. anthracis has been reported recently.130 This molecular assay is based on the combination of molecular beacons and real-time PCR technologies. Molecular beacons are highly specific nucleic acid probes that, when bound to their target DNA sequences, act as switches by emitting fluorescence as a result of the alteration of their conformation.130,131 The molecular beacon– based real-time PCR assay was performed by designing five molecular beacons targeting B. anthracis capA, capB, capC, lef, and pag alleles that were used in five uniplex assays for the detection of these genes in a broad range of samples.130 The assay is conducted in 25 μL reaction mixture containing 5.0 μL DNA, 4.5 μL distilled water, 12.5 μL Platinum Quantitative PCR SuperMix-UDG, 1.0 μL of one of the five

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Bacillus

molecular beacons and 1.0 μL (20 pmol/μL) of each of the forward and reverse primers. The assay is run on a 7900HT Real-Time PCR System (Applied Biosystems) with the following thermal cycling program: denaturation (94°C for 10 min), followed by 40 cycles of amplification (94°C denaturation for 15 s; 50°C annealing and data collection for 30 s; and 72°C polymerization for 30 s). During each cycle of amplification, the fluorescence emission in each sample is automatically recorded at 490 nm. Another innovative powerful gene-amplification technique that is becoming very popular and emerging as a simple and rapid diagnostic tool for the detection and identification of microbial diseases is the loop-mediated isothermal amplification (LAMP) that was described for the first time by Notomi et al.132 LAMP is based on the principle of strand-displacing DNA synthesis by the Bst DNA polymerase, with six distinct primers that recognize eight distinct sequences of a target gene. DNA amplification performed under isothermal conditions (60°C–65°C) using a heating block and/or water bath, is usually completed in less than 1 h. The gene-amplification products can be analyzed by agarose gel electrophoresis or by real-time monitoring of the turbidity of the reaction mixture in a turbidimeter. In this later case, it is also possible to quantify the gene copy number based on a standard curve generated from different concentrations of the gene copy number plotted against time of positivity. Moreover, gene amplification can be monitored by the naked eye either as turbidity or in the form of a color change when a fluorescent dsDNA intercalating dye is used. LAMP assays have been used for rapid detection of several pathogenic viruses, bacteria, and blood protozoa.133,134 Recently, a specific LAMP assay targeting the chromosomal sap gene that codifies for the S-layer constituent protein and cap and pag plasmid genes was designed by Kurosaki et  al.135 as a method for B. anthracis detection. Loopamp DNA Amplification Kit (Eiken Chemical, Tokyo, Japan) is used for performing LAMP reaction following the manufacturer’s instructions. Briefly, a 25 µL final volume reaction mixture containing 40 pmol of each forward inner primer (FIP) and a reverse inner primer (BIP); 5 pmol of each of the outer primers F3 and B3; 20 pmol of each of the two loop primers (LF and LB); 12.5 µL of 2× reaction mix; 1.0 µL of Bst DNA polymerase; and 5.0 µL of DNA sample are incubated at 63°C for 60 min. The LAMP products are analyzed on 1% agarose gel and ehidium bromide staining. Otherwise, for real-time monitoring of amplification, the reaction mixtures are incubated and observed by spectrophotometric analysis using a real-time turbidimeter (LA-200; Teramecs, Kyoto, Japan). The time of positivity observed through realtime LAMP assay is determined as the time at which the turbidity reached the threshold value fixed at 0.1, which is double the average turbidity value of negative controls in several replicates. The authors reported that this assay is 10- to 100-fold more sensitive than conventional PCR-based methods, allowing the detection of 10 fg of bacterial DNA per reaction and 3.6 CFU of bacterial spores in blood samples from experimentally infected mice.

21.3  C  ONCLUSIONS AND FUTURE PERSPECTIVES Spores produced by Bacillus spp., other than B. anthracis, are ubiquitous in the environment owing to their resistance to harsh environmental conditions as well as to disinfection procedures. Their presence in food samples or clinical specimens has often been considered as a contamination. However, Bacillus spp. can cause serious and even lethal infections or intoxications not only in immunocompromised individuals or patients with other high-risk factors but also in otherwise healthy individuals. Conventional diagnostic procedures are based mainly on phenotypic and biochemical characteristics. For example, in the B. cereus group, the differentiation of the well-known opportunistic human pathogen B. cereus and the anthrax agent B. anthracis is based in part on the presence of a poly-gamma-D-glutamic acid capsule. However, both encapsulated B. cereus that has been implicated in fatal pneumonia and B. thuringiensis strains producing a capsule similar to the B. anthracis one136 have been discovered recently. This finding underlines the importance of using multiple tests for a reliable identification of isolates associated with human disease. Different Bacillus spp. outbreaks in hospital environments where the origin of the disseminated isolate was the reusable equipment, such as towels, gowns, and hand ventilator balloons, have been reported. Also, Bacillus spp. have been isolated from alcoholic cotton and detergents. These reported cases highlight the importance of using disposal or wrapping steamed towels and wrapping alcoholic cotton as preventive measures to avoid nosocomial bacteremia caused by these spore formers. They also underscore the importance of using professionally performed sterilization procedures for all reusable equipment and that should include eradication of spores. For this purpose, autoclaving is imperative and especially for reusable equipment in intensive care units and for material used by immunocompromised patients. The 70%–90% alcohol that is extensively used in hospitals is suitable as germicide but does not affect spores. On the other hand, regular cleaning of the filters of the air-conditioning system and the inlets and outlets of the air-ventilation system, as well as using the recommended concentration of sodium hypochlorite in the laundry process would be important measures to avoid nosocomial bacteremia by spore formers. Several molecular detection methods are actually available for B. anthracis and its pathogenecity factors, including the secreted anthrax toxins and the capsule, as well as for B.  cereus and its toxins. These include conventional PCR, multiplex PCR and RT–PCR. However, although highly specific and sensitive, these molecular methods have some drawbacks. The conventional PCR bears the risk of false-positive results due to sample contamination. The multiplex PCR could be a good alternative for detecting different pathogens or different toxins of the same pathogenic bacterial species simultaneously. However, in some cases, it is possible to get false-negative results because of the presence of different primer pairs in the same reaction mix or different annealing

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temperatures. In addition, this method is qualitative and does not allow quantification of the pathogenic bacterium or of its toxins. In order to quantify the presence of a specific pathogenic bacterium or its toxins, TaqMan or SYBR greenbased RT–PCRs could be applied. However, although highly sensitive, this technique could be biased by the presence of PCR inhibitors in clinical specimens. Also, the presence of dominant concentrations of human DNA in comparison to the DNA from the pathogenic bacterium in samples such as blood and cerebrospinal fluid, in particular, during the first step of the infection, can deeply affect the sensitivity of the PCR. Another practical drawback of RT–PCR is the necessity of expensive and sophisticated real-time platforms and reagents that are not available in most laboratories especially in developing countries. During this last decade, and since its description in 2000, LAMP technique has been largely used for the diagnosis of human pathogenic bacteria as well as viruses and protozoa. Only one recent report described the application of this assay for the diagnosis of B. anthracis in experimentally infected mice. The advantages of this technique consist of rapid amplification at one set temperature without the need of a thermal cycler, simple operation, and easy detection by a turbidimeter or naked eye. Also, combining LAMP assay with boiling extraction, it is possible to quickly detect very low copies of the target gene and hence to perform an efficient on-site diagnosis in less than 1 h without the need for culture growth or precise DNA extraction. Hence, LAMP can be considered an innovative technique that has potential applications for both clinical diagnosis and surveillance of infectious diseases in developing countries, without requiring sophisticated equipment or skilled personnel. It is, however, important to take into account that one of the major drawbacks of this assay is that designing the primers presents a difficulty because at least six distinct primers that recognize eight distinct sequences of a target gene are required for the performance of the assay. In conclusion, the most serious and important human pathogen within the genus Bacillus is B. anthracis. However, other species should also be considered in the diagnosis of septicemic infections, pneumonia, meningitis, endophthalmitis, and cutaneous infections. Among others, Bacillus cereus is being acknowledged as an emerging pathogen, especially in immunocompromised individuals. Also, although rare, capsule-producing pathogenic B. cereus strains with fatal anthrax-like pathology exist and should be considered, especially in patients with high-risk occupational factors, such as metal workers, and in particular, in zones where anthrax was endemic. As far as the detection of Bacillus spp. in clinical specimens, a combination of culture-based and molecular methods, including PCR amplification of the virulence factor encoding genes is necessary for the confirmation of the diagnosis and hence the choice of the appropriate therapy. To avoid misdiagnosis and unnecessary treatment, performing more than one set of cultures from clinical specimens is recommended before assuming that an isolate is a pathogen or dismissing a pathogen as a contaminant, which can

Molecular Detection of Human Bacterial Pathogens

cause complications and even mortality due to delay in the treatment.71

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228 63. van Sorge, N.M. et al., Anthrax toxins inhibit neutrophil signaling pathways in brain endothelium and contribute to the  pathogenesis of meningitis, PLoS One, 3, doi:10.1371/ journal.pone.0002964, 2008. 64. Sejvar, J.J., Tenover, F.C., and Stephens, D.S. Management of anthrax meningitis, Lancet. Infect. Dis., 5, 287, 2005. 65. Simonson, T.S. et al., Bacillus anthracis in China and its relationship to worldwide lineages, BMC Microbiol., 9, 71, 2009. 66. Doganay, M., and Metan, G., Human Anthrax in Turkey from 1990 to 2007, Vector-born. Zoonot., 9, DOI: 10.1089/ vbz.2008.0032, 2009. 67. Lanska, D., Anthrax meningoencephalitis, Neurology, 59, 327, 2002. 68. Narayan, S.K. et al., Anthrax meningoencephalitis—declining trends in an uncommon but catastrophic CNS infection in rural Tamil Nadu, South India, J. Neurol. Sci., 281, 41, 2009. 69. Weber, D.J. et al., Clinical significance of Bacillus species isolated from blood cultures, South. Med. J., 82, 705, 1989. 70. Lee, C.C. et al., Clinical significance of potential contaminants in blood cultures among patients in a medical center, J. Microbiol. Immunol. Infect., 40, 438, 2007. 71. Ashkenazi-Hoffnung, L. et al., Seasonality of Bacillus species isolated from blood cultures and its potential implications, Am. J. Infect. Control, 37, 495, 2009. 72. Van Der Zwet, W.C. et al., Outbreak of Bacillus cereus infections in a neonatal intensive care unit traced to balloons used in manual ventilation, J. Clin. Microbiol., 38, 4131, 2000. 73. Hernaiz, C. et al., Nosocomial bacteremia and catheter infection by Bacillus cereus in an immunocompetent patient, Clin. Microbiol. Infect., 9, 973, 2003. 74. Ghelardi, E. et al., Bacillus thuringiensis pulmonary infection: Critical role for bacterial membrane-damaging toxins and host neutrophils, Microbes Infect., 9, 591, 2007. 75. Ozkocaman, V. et al., Bacillus spp. among hospitalized patients with haematological malignancies: Clinical features, epidemics and outcomes, J. Hosp. Infect., 64, 169, 2006. 76. Lépine, A. et al., Bacillus licheniformis septicemia in a verylow-birth-weight neonate: A case report, Infection, 37, 156, 2009. 77. Blue, S.R., Singh, V.R., and Sanbolle, M.A., Bacillus licheniformis bacteraemia: Five cases associated with indwelling central venous catheters, Clin. Infect. Dis., 20, 629, 1995. 78. Haymore, B.R. et al., A case of persistent Bacillus pumilis bacteremia associated with cholangitis, J. Infect., 52, 154, 2006. 79. Bentur, H.N., Dalzell, A.M., and Riordan, F.A.I. Central venous catheter infection with Bacillus pumilus in an immunocompetent child: A case report, Ann. Clin. Microbiol. Antimicrob., 6, 12, 2007. 80. Ramakrishnan, R. et al., Microbiological profile of culture proven cases of exogenous and endogenous endophthalmitis: A 10-year retrospective study, Eye, 23, 945, 2009. 81. Ramos-Esteban, J.C. et al., Bacillus megaterium delayed onset lamellar keratitis after LASIK, J. Refract. Surg., 22, 309, 2006. 82. Callegan, M.C. et al., Virulence factor profiles and antimicrobial susceptibilities of ocular bacillus isolates, Curr. Eye. Res., 31, 693, 2006. 83. Kopel, A.C., Carvounis, P.E., and Holz E.R. Bacillus cereus endophthalmitis following intravitreous bevacizumab injection, Ophthalmic. Surg. Lasers. Imaging., 39, 153, 2008. 84. Callegan, M.C. et al., Bacillus endophthalmitis: Roles of bacterial toxins and motility during infection, Invest. Ophthalmol. Vis. Sci., 46, 3233, 2005.

Molecular Detection of Human Bacterial Pathogens 85. Ramadan, R.T. et al., A role for tumor necrosis factor-alpha (TNFα) in experimental Bacillus cereus endophthalmitis pathogenesis, Invest. Ophthalmol. Vis. Sci., 49, 4482, 2008. 86. Moyer, A.L. et al., Bacillus cereus-induced permeability of the blood-ocular barrier during experimental endophthalmitis, Invest. Ophthalmol. Vis. Sci., 50, 3783, 2009. 87. Mahler, H. et al., Fulminant liver failure in association with the emetic toxin of Bacillus cereus, N. Engl. J. Med., 336, 1142, 1997. 88. Latsios, G. et al., Liver abscess due to Bacillus cereus: A case report, Clin. Microbiol. Infect., 9, 1234, 2003. 89. Hoffmaster, A.R. et al., Identification of anthrax toxin genes in a Bacillus cereus associated with an illness resembling inhalation anthrax, Proc. Natl. Acad. Sci. USA, 101, 8449, 2004. 90. Miller, J.M. et al., Fulminating bacteremia and pneumonia due to B. cereus, J. Clin. Microbiol., 35, 504, 1997. 91. Avashia, S.B. et al., Fatal pneumonia among metalworkers due to inhalation exposure to Bacillus cereus containing Bacillus anthracis toxin genes, Clin. Infect. Dis., 44, 414, 2007. 92. Hoffmaster, A.R. et al., Characterization of Bacillus cereus isolates associated with fatal pneumonias: strains are closely related to Bacillus anthracis and harbor B. anthracis virulence genes, J. Clin. Microbiol., 44, 3352, 2006. 93. Katsuya, H. et al., A patient with acute myeloid leukemia who developed fatal pneumonia caused by carbapenem-resistant Bacillus cereus, J. Infect. Chemother., 15, 39, 2009. 94. Helgason, E. et al., Genetic structure of population of Bacillus cereus and B. thuringiensis isolates associated with periodontitis and other human infections, J. Clin. Microbiol., 38, 1615, 2000. 95. Johnson, B.T. et al., Extracellular proteolytic activities expressed by Bacillus pumilus isolated from endodontic and periodontal lesions, J. Med. Microbiol., 57, 643, 2008. 96. Yamane, K. et al., Identification and characterization of clinically isolated biofilm-forming gram-positive rods from teeth associated with persistent apical periodontitis, J. Endod., 35, 347, 2009. 97. Sunde, P.T. et al., Microbiota of periapical lesions refractory to endodontic therapy, J. Endod., 28, 304, 2002. 98. Ewald C., Kuhn, S., and Kalff, R. Pyogenic infections of the central nervous system secondary to dental affection—a report of six cases, Neurosurg. Rev., 29, 163, 2006. 99. Lequin, M.H. et al., Bacillus cereus meningoencephalitis in preterm infants: Neuroimaging characteristics, Am. J. Neuroradiol., 26, 2137, 2005. 100. Manickam, N., Knorr, A., and Muldrew, K.L. Neonatal meningoencephalitis caused by Bacillus cereus, Pediatr. Infect. Dis. J., 27, 843, 2008. 101. Evreux, F. et al., A case of fatal neonatal Bacillus cereus meningitis, Arch. Pediatr., 14, 365, 2007. 102. Lebessi, E. et al., Bacillus cereus meningitis in a term neonate, J. Matern. Fetal. Neonatal. Med., 22, 458, 2009. 103. de Almeida, S.M. et al., Fatal Bacillus cereus meningitis without inflammatory reaction in cerebral spinal fluid after bone marrow transplantation, Transplantation, 76, 1533, 2003. 104. Haase, R. et al., Successful treatment of Bacillus cereus meningitis following allogenic stem Cell transplantation, Pediatr. Transplant., 9, 338, 2005. 105. Meredith, F.T. et al., Bacillus cereus necrotizing cellulitis mimicking clostridial myonecrosis: Case report and review of the literature, Scand. J. Infect. Dis., 29, 528, 1997. 106. Sada, A., Necrotizing fasciitis and myonecrosis “synergistic necrotizing cellulitis” caused by Bacillus cereus, J. Dermatol., 36, 423, 2009.

Bacillus 107. Pinedo, S., Bos, A.J., and Siegert, C.E. Relapsing Bacillus cereus peritonitis in two patients on peritoneal dialysis, Perit. Dial. Int., 22, 424, 2002. 108. Monteverde, M.L. et al., Relapsing Bacillus cereus peritonitis in a pediatric patient on chronic peritoneal dialysis, Perit. Dial. Int., 26, 715, 2006. 109. Berry, N. et al., Bacillus circulans peritonitis in a patient treated with CAPD, Perit. Dial. Int., 24, 488, 2004. 110. Park, D.J. et al., Relapsing Bacillus licheniformis peritonitis in a continuous ambulatory peritoneal dialysis patient, Nephrology, 11, 21, 2006. 111. Shimoni, Z. et al., Bacillus cereus peritonitis after Cesarean section, Surg. Infect., 9, 105, 2008. 112. Knisely, R.F., Selective medium for Bacillus anthracis, J. Bacteriol., 92, 784, 1966. 113. Juergensmeyer, M.A. et al., A selective chromogenic agar that distinguishes Bacillus anthracis from Bacillus cereus and Bacillus thuringiensis, J. Food. Protect., 69, 2002, 2006. 114. Marston, C.K. et al., Evaluation of two selective media for the isolation of Bacillus anthracis, Lett. Appl. Microbiol., 47, 25, 2008.‌ 115. Klee, S.R. et al., Characterization of Bacillus anthracis-like bacteria isolated from wild great apes from Cote d’Ivoire and Cameroon, J. Bacteriol., 188, 5333, 2006. 116. Hao, R. et al., Rapid detection of Bacillus anthracis using monoclonal antibody functionalized QCM sensor, Biosens. Bioelectron. 24, 1330, 2009. 117. Tang, S. et al., Detection of anthrax toxin by an ultrasensitive immunoassay using europium nanoparticles, Clin. Vaccine. Immunol., 16, 408, 2009. 118. Duriez, E. et al., Femtomolar detection of the anthrax edema factor in human and animal plasma, Anal. Chem., 81, 5935, 2009. 119. AuCoin, D.P. et al., Rapid detection of the poly-γ-D-glutamic acid capsular antigen of Bacillus anthracis by latex agglutination, Diagn. Microbiol. Infect. Dis., 64, 229, 2009. 120. Schofield, D.A., and Westwater, C., Phage-mediated bioluminescent detection of Bacillus anthracis, J. Appl. Microbiol., 107, 1468, 2009. ‌ 121. Dauphin, L.A., Moser, B.D., and Bowen, M.D. Evaluation of five commercial nucleic acid extraction kits for their ability to inactivate Bacillus anthracis spores and comparison of DNA yields from spores and spiked environmental samples, J. Microbiol. Methods, 76, 30, 2009. 122. Dauphin, L.A. et al., Gamma irradiation can be used to inactivate Bacillus anthracis spores without compromising the

229 sensitivity of diagnostic assays, Appl. Environ. Microbiol., 74, 4427, 2008. 123. Dauphin, L.A., and Bowen, M.D., A simple method for the rapid removal of Bacillus anthracis spores from DNA preparations, J. Microbiol. Methods, 76, 212, 2009. 124. Sambrook, J., Fritsch, E.F., and Maniatis, T., Molecular Cloning: A Laboratory Manual, 2nd ed., Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY, 1989. 125. Horz, H.P. et al., Selective isolation of bacterial DNA from human clinical specimens, J. Microbiol. Methods, 72, 98, 2008. 126. Hansen, W.L., Bruggeman, C.A., and Wolffs, P.F. Evaluation of new preanalysis sample treatment tools and DNA isolation protocols to improve bacterial pathogen detection in whole blood, J. Clin. Microbiol., 47, 2629, 2009. 127. Ramisse, V. et al., The Ba813 chromosomal DNA sequence effectively traces the whole Bacillus anthracis community, J. Appl. Microbiol., 87, 224, 1999. 128. Qi, Y. et al., Utilization of the rpoB gene as a specific chromosomal marker for real-time PCR detection of Bacillus anthracis, Appl. Environ. Microbiol., 67, 3720, 2001. 129. Antwerpen, M.H. et al., Real-time PCR system targeting a chromosomal marker specific for Bacillus anthracis, Mol. Cell. Probes, 22, 313, 2008. 130. Hadjinicolaou, A.V., and Victoria, L., Use of molecular beacons and multi-allelic real-time PCR for detection of and discrimination between virulent Bacillus anthracis and other Bacillus isolates, J. Microbiol. Methods, 78, 45, 2009. 131. Li, Y. et al., Molecular beacons: An optimal multifunctional biological probe, Biochem. Biophys. Res. Comm., 373, 457, 2008. 132. Notomi, T. et al., Loop-mediated isothermal amplification of DNA, Nucleic Acids Res., 28, E63, 2000. 133. Mori, Y. et al., Detection of loop-mediated isothermal amplification reaction by turbidity derived from magnesium pyrophosphate formation, Biochem. Biophys. Res. Comm., 289, 150, 2001. 134. Parida, M. et al., Loop mediated isothermal amplification (LAMP): A new generation of innovative gene amplification technique; perspectives in clinical diagnosis of infectious diseases, Rev. Med. Virol., 18, 407, 2008. 135. Kurosaki, Y. et al., A simple and sensitive method for detection of Bacillus anthracis by loop-mediated isothermal amplification, J. Appl. Microbiol., 107, 1947, 2009. ‌ 136. Cachat, E. et al., A Bacillus thuringiensis strain producing a polyglutamate capsule resembling that of Bacillus anthracis, FEMS Microbiol. Lett., 285, 220, 2008.

22 Enterococcus Teresa Semedo-Lemsaddek, Paula Lopes Alves, Rogério Tenreiro, and Maria Teresa Barreto Crespo CONTENTS 22.1 Introduction...................................................................................................................................................................... 231 22.1.1 Classification and Morphology............................................................................................................................. 231 22.1.2 Clinical Features, Epidemiology, and Pathogenesis............................................................................................. 231 22.1.3 Diagnosis.............................................................................................................................................................. 233 22.1.3.1 Phenotypic Techniques.......................................................................................................................... 233 22.1.3.2 Molecular Techniques............................................................................................................................ 233 22.2 Methods............................................................................................................................................................................ 241 22.2.1 Sample Preparation............................................................................................................................................... 241 22.2.2 Detection Procedures............................................................................................................................................ 241 22.2.2.1 Semiautomatic and Automatic Identification......................................................................................... 241 22.2.2.2 Multiplex PCR for the Detection of Enterococcus Species................................................................... 242 22.3 Conclusions and Future Perspectives............................................................................................................................... 243 Acknowledgments...................................................................................................................................................................... 243 References.................................................................................................................................................................................. 243

22.1  INTRODUCTION 22.1.1  Classification and Morphology Enterococci are mainly viewed as commensal bacteria inhabiting the gastrointestinal tract of humans and animals, but in fact, these microorganisms have been recognized as potentially pathogenic for humans for more than a century. In Thiercelin’s1 original description, these bacteria had been isolated from patients with enteritis, appendicitis, and meningitis.2 The classification and nomenclature of the genus Enterococcus has raised interest and constant discussion over the years, ever since Thiercelin first proposed the name “enterocoques” in 1902. In 1970, Kalina3 proposed that Streptococcus faecalis and S. faecium should be renamed as Enterococcus faecalis and E. faecium, respectively. These proposals were not officially recognized for many years, and the genus Enterococcus did not appear in the Approved List of Bacterial Names until the 1980s. In the 1980s, Schleifer and Kilpper-Bälz4 resurrected the genus Enterococcus after performing DNA–DNA and DNA–rRNA hybridizations. Other studies carried out in the same decade, including 16S rRNA sequencing,5 also demonstrated that enterococci form a separate genus. Analysis of these molecular data divided the streptococci sensu lato into three genera: (i) Streptococcus sensu stricto, comprising the majority of the known species; (ii) Enterococcus,

for the enteric group; and (iii) Lactococcus, for the lactic streptococci. Since then, the genus Enterococcus has been accepted as valid, and today it comprises 33 species. In 2007, Kohler6 performed an extensive review of the publications that lead to each of the species currently accepted and included in TOBA (Taxonomic Outline of the Bacteria and Archaea),7 based on a comprehensive taxonomy using 16S rRNA gene phylogeny. Members of the genus Enterococcus are gram-positive ovoid cells that appear singly, in pairs, or in short chains; are catalase- and oxidase-negative; facultative anaerobes; and homofermentative, with lactic acid as the end product of glucose fermentation. The majority of the enterococci grow from 10ºC to 45ºC, with 6.5% NaCl and at pH 9.6, hydrolyze esculin in the presence of 40% bile salts and the G + C content ranges from 37% to 45%.8,9 However, not all enterococci exhibit these characteristics; several authors have analyzed and reported differences in physiological properties among different species and strains belonging to the same species,10–15 making a reliable identification at species level difficult.

22.1.2  Clinical Features, Epidemiology, and Pathogenesis Although enterococci have traditionally been regarded as low-grade pathogens, in the last decades they have emerged 231

232

as an increasingly important cause of nosocomial infections, mainly in patients with underlying disease. All kinds of severe immunosupression (e.g., oncology, hematology, nephrology, or transplantation units), long hospital stays,16 or the presence of urinary or vascular catheters constitute notable risk factors.17 The infections most frequently caused by enterococci are urinary tract infections (UTIs),18 followed by intraabdominal/ intrapelvic abscesses or postsurgery wound and bloodstream infections (BSIs).19 Enterococci are also infrequent causes of central nervous system (CNS) infections, neonatal infections, respiratory tract infections, osteomyelitis, or cellulitis.20 The importance of enterococci in the hospital setting is increasing every year. Enterococci are the third most common cause of nosocomial blood infections in the United States,21 and in Europe, the fourth (http://www.earss.rivm. nl/). Regarding urinary tract infections, they rank second in both the United States and Europe.22,23 Although more than thirty species have been identified,7 only two are responsible for the majority of human infections. The species E. faecalis and E. faecium account for 80% and 20% of the enterococcal infections, respectively. The proportion of E. faecium infections is increasing, and other species are being more frequently isolated from clinical samples, namely, E. gallinarum, E. casseliflavus, and E. raffinosus.22,24,25 The inherent tolerance of enterococci to unfavorable conditions allows them to survive for long periods in hospital settings, even on dry surfaces, and medical staff and instruments can be responsible for the transmission between hospitalized patients.17,26–28 These characteristics contribute to an increased risk of enterococcal nosocomial infections and emphasize the importance of bacterial control in the health care environment. Enterococcal infections are an important health concern, and as nosocomial pathogens, they have a direct and significant economic impact by prolonging hospital stays and requiring additional therapeutic treatments.29 Another important factor contributing to the increasing number of enterococcal infections is the uncontrolled use of antimicrobial agents, which enhances the selective pressure for bacteria with natural and/or acquired antibiotic resistances, such as enterococci. Administration of antimicrobial agents that have little effect on enterococci results in the elimination of many endogenous microbiota and may open new niches for the overgrowth of pathogenic strains. Enterococci have intrinsic resistance to several antimicrobial agents (e.g., cephalosporins, lincosamides, many β-lactams and low levels of aminoglycosides); frequently acquire plasmids and transposons carrying antimicrobial resistance genes (e.g., chloramphenicol, erythromycin, tetracyclin, and glycopeptides such as vancomycin) that can subsequently be transferred to other strains sharing the same ecological niche.2,16,26,28 Vancomycin resistance gains increased importance because this antimicrobial is considered a last resource treatment for many human infections. Vancomycinresistant enterococci (VRE) were first detected in Europe

Molecular Detection of Human Bacterial Pathogens

(United  Kingdom, and France) in 1988, and in the following year in the United States.30 In the last decades, the incidence of VRE rose, mainly due to E. faecium isolates, and vancomycin-resistant strains have already been reported in countries throughout the world, such as Australia, Belgium, Canada, Denmark, Germany, Italy, Malaysia, Netherlands, Spain, Sweden, and Portugal.31 While the increase of VRE in the United States is attributed to the uncontrolled usage of antimicrobials in hospitals,31 in Europe the reservoir of resistant strains seems to be linked with the use of avoparcin as a feed supplement for various livestock, and contamination via the food chain has been suggested.32–35 The usage of pulsed field gel electrophoresis has also demonstrated that acute outbreaks are typically related to a single VRE clone or a limited number of clones.36–38 Asymptomatic patients with VRE colonizing the gastrointestinal tract are usually the reservoir, and the dissemination between patients is mainly attributed to the indirect contact of the hands of health care workers. Surprisingly, little is known about the factors that contribute to the ability of enterococci to cause infection. In E. faecalis, several putative virulence factors have been described over the last years, involved either in the attachment to host cells or to extracellular matrix proteins, or implicated in cell and tissue damage. For gastrointestinal commensals such as enterococci, adhesins that promote binding to host cell receptors are expected to play an important role in the establishment and maintenance of colonization.39 The enterococcal surface protein (Esp) is a high molecular weight extracellular surface protein of unknown function whose frequency is increased among infection-derived isolates.40 This protein contributes to colonization and persistence of enterococci in the urinary tract41 and also to biofilm formation on abiotic surfaces.42 Aggregation substance (AS) is a pheromone-inducible surface protein that mediates the binding of donor cells to plasmid free recipients.43,44 This protein is essential for high-efficiency conjugation of sex pheromone plasmids, contributing to the dissemination of antimicrobial resistance and virulence determinants. Different functions have been attributed to AS in addition to bacterial cell aggregation, including adherence to host tissues45,46 and internalization,47 adhesion to fibrin and increased cell-surface hydrophobicity,48 resistance to killing by polymorphonuclear leukocytes and macrophages,46,49 and increased vegetation size in experimental endocarditis50 and endophthalmitis models.51 Another adhesion-associated protein was identified while analyzing serum from patients with E. faecalis endocarditis (named EfaA for E. faecalis antigen A).52,53 Regarding cell and tissue damage, the first virulence factor to be studied was cytolysin, a posttranslationally modified protein toxin that causes a β-hemolytic reaction on certain blood erythrocytes and also possesses bactericidal activity against a broad range of gram-positive bacteria.54–56 Gelatinase is an extracellular zinc-endopeptidase capable of hydrolysing gelatin, collagen, casein, and other small biologically active peptides, and its potential contribution to virulence has been suggested in several studies.57–60

Enterococcus

As regards E. faecium, only three putative virulence genes have been identified; they encode an enterococcal surface protein (espefm), a putative hyaluronidase (hyl)61,62 and an endocarditis-related adhesin EfaAfm.52,53 The putative virulence factors described for the genus Enterococcus can either be encoded by mobile genetic elements, such as plasmids or transposons, or by the bacterial chromosome. These genomic region clustering genes that contribute, directly or indirectly, to the pathogenic potency of the harboring microorganism are termed pathogenicity islands or PAIs.63 The generalized sequencing of genomes revealed that PAIs are much more widespread than previously thought.63 Regarding the genus Enterococcus, two PAIs have been described: one in a vancomycin-resistant strain of E. faecalis that caused a nosocomial outbreak,64 and the other in a group of nosocomial strains of E. faecium.65 More recently, the worldwide dissemination of the E. faecalis PAI among isolates from diverse locations and origins was demonstrated.66

22.1.3  Diagnosis 22.1.3.1  Phenotypic Techniques Detection and reliable identification, especially regarding VRE, is of major concern since VRE leaves very few options for the treatment of enterococcal infections.16,28 At the present time in Europe, VRE infections are increasing; meanwhile, in the United States, colonization of hospitalized patients is endemic in many tertiary care hospitals and increasingly causes infections.16 Conventional culture methods remain the most reliable and accurate techniques for the detection of bacterial pathogens such as enterococci. Traditionally, enterococcal identification relied on various cultivation procedures associated with the analysis of phenotypic profiles including Gram staining, colony morphology, motility, pigmentation, growth requirements, and enzymatic and/or metabolic activities. However, this culture-based identification is time consuming and frequently limited by the variability of atypical strains, the lack of sufficient data on recently described species, and the low reproducibility of some tests. Since Enterococcus is neither a phylogenetically nor phenotypically coherent and homogeneous genus, manual or automated systems based on phenotypic characteristics do not always allow the correct strain identification; therefore these problems assume great importance.9 Another disadvantage of the standard culture methods is their lack of ability to detect a substantial proportion of the bacterial population that is nonculturable. These viable but nonculturable (VBNC) microorganisms demonstrate metabolic activity,67,68 maintain their pathogenicity69,70 and their antibiotic resistance traits,71 can express72 and exchange genes,73 and are able to reestablish division upon restoration of favorable conditions.74,75 It has been previously demonstrated that E. faecalis can enter the VBNC state,72 thus representing a risk in hospital settings and emphasizing the importance of molecular

233

detection in substitution of standard culture methods. Consequently, the optimization of molecular methods for microbial analysis is becoming more important every day, with special relevance for rapid, easy-to-implement, and reliable procedures that may be applied in routine laboratories. 22.1.3.2  Molecular Techniques Nucleic acid–based identification and detection methods have been developed for nearly all bacterial pathogens.76 Rapid and sensitive DNA-based assays that are applicable for the direct detection of enterococci in human clinical samples may improve the quickness and the accuracy of the detection of these bacteria. In spite of all the advantages associated with molecularbased methodologies, the detection of microorganisms in clinical samples can come across several problems, such as PCR inhibition by sample-associated components, crosscontamination in sensitive PCR assays, and detection of living as well as dead cells (this specific problem can be solved by using mRNA as amplification target instead of DNA in RT–PCR). Although PCR amplification of species-specific targets enables precise identification of known microbial species with short reporting times, it is also important to highlight that positive amplification only demonstrates the presence of the appropriate target DNA sequences in the sample and does not imply that the target organism is viable. This problem can be overcome by the evaluation of cell viability (mRNA detection by RT–PCR) or visualization of living cells in mixed bacterial populations (in situ PCR/hybridization). Several molecular diagnostic methods based on the detection of bacterial nucleic acids have already been described for enterococci. The majority of these methodologies relies on PCR due to the low amounts of DNA and/or cells needed, as well as the detection of a specific DNA sequence against a large background of prokaryotic and eukaryotic cells and organic material present in mixed samples.77 Under ideal conditions, with 100% efficiency and unlimited resources, a single copy of target gene can be amplified to approximately 3.4 × 1010 copies after 35 cycles of PCR. However, it becomes important to emphasize that the ability to design oligonucleotide primers is limited by our knowledge of a microorganism’s genome, as well as by the representativeness of publicly available sequence databases in terms of all variants of that microbial group. Distinct molecular approaches have already been described for enterococci including broad-range PCR,78 multiplex PCR,79– 81 species-specific PCR,82–87 real-time PCR,76,88,89 direct detection of bacteria using fluorescent in situ hybridization (FISH) probes,90–92 and amplification of 16S and 23S ribosomal gene sequences for hybridization to a microarray.93,94 Conventional PCR assays detect only the presence or absence, rather than the quantity, of a target microorganism and cannot distinguish between nonviable and viable cells. To deal with these limitations several modifications of standard PCR have recently been developed.

234

Real-time quantitative PCR (qPCR), such as the TaqMan system, relies on the release and detection of a signal after cleavage of a fluorescent-labeled probe by the 5′–3′-exonuclease activity of Taq DNA polymerase. A fluorogenic oligonucleotide probe, which specifically binds within the two PCR primer regions, is degraded in each PCR cycle, thereby monitoring the amplification of the target gene by an increasing fluorescence signals in real time. The signal is related to the amount of amplicon present during each cycle and will increase as the amount of specific amplicon increases.76,88,89,95,96 The specificity of the probe introduces a high level of confidence concerning the detection of target microorganisms in mixed samples. Another approach with known advantages is hybridization of whole bacterial cells with fluorescent oligonucleotides (FISH) targeted to broad-range sequences (16 or 23S rRNA) or specific genes.91–93 This method is rapid and easy-to-perform and allows the in situ observation of the abundance, spatial distribution, and morphology of bacterial cells.92 FISH assays can significantly shorten time to retrieve results and to initiate control measures in the food industry. Microarray technology is also a powerful tool that can be used for simultaneous detection of large sets of different genes of DNA (or RNA) targets on a glass slide.76 By processing multiple samples and analyzing up to tens of thousands of genes in a single assay, this method is a promising alternative for monitoring environmental and food bacteria, but it requires expensive microarray and laser-scanner apparatus. When applying microarrays for microorganism detection, PCR amplification of target genes is often involved in order to enhance detection sensitivity.76,97–99 Microarray methodology has already been applied for the detection of enterococci in environments such as wastewater76 and artificially contaminated milk samples.94 A summary of available probes for the most relevant enterococcal human pathogens that can be used in qPCR, RT–qPCR, hybridization assays, FISH, and microarrays is presented in Table 22.1. As stated before, a variety of molecular methods have been developed for enterococcal detection and identification at the genus or species level. The detected DNA targets include genes encoding for rRNA,76,84,88,89,103,104 the alpha subunit of RNA polymerase and the phenylalanyl-tRNA synthase,106 the elongation factor EF-Tu,83 the d-alanine:dalanine ligase,80 a penicillin-binding protein,77 the muramidase,87 the alpha subunit of the bacterial ATP synthase,107 the manganese-dependent superoxide dismutase,82,108 and the heat-shock proteins (groESL),76,86,109,110 as well as genus or species-specific antimicrobial determinants.80,81,89 A short description of each method is presented below and additional information about broad-range PCR primers (designed for eubacteria and lactic acid bacteria and to be used in positive control reactions) and enterococcal PCR primers (designed for selective amplification at genus and species level) can be found in Table 22.2. Regarding specific primers for identification at species level, Table 22.2 only includes the most relevant human pathogens (E. faecalis, E. faecium,

Molecular Detection of Human Bacterial Pathogens

E. casseliflavus/E. flavescens, and E. gallinarum) and for information about other enterococcal species, see SemedoLemsaddek et al.140 Ribosomal RNA. rRNA is a universal component of bacterial ribosomes and 5S, 16S, and 23S rRNAs are present at the small (30S) or large (50S) subunits that comprise the complete active ribosome (70S). Since rRNAs are found in high copy numbers (103–104 molecules) per actively growing cell and their coding genes are usually present in more than two copies, targeting the rRNAs or their coding genes has the potential to increase the detection sensitivity compared to assays based on the detection of single-copy genes.121 Small subunit ribosomal RNA (16S rRNA) gene sequences are widely used as a target for the development of PCR primers and/or specific hybridization probes due to their phylogenetic informational content and wide availability in databases. By comparison, although the large subunit rRNA (23S rRNA) gene contains more sequence variations than the 16S rRNA gene, and thus may provide more useful targets for microorganism identification at species or intraspecies level, it has been less frequently used, mainly because of the limited availability of sequence information. Since both 16S rRNA and 23S rRNA are part of the ribosome, and, due to evolutionary constraints, there is a minimal variation within these regions. So, examining the 16S-23S rRNA intergenic noncoding region (ITS) may be necessary to distinguish very similar microorganisms. Because of the universality of rRNA genes, the selection of specific primers or probes has to be criteriously performed. The use of such gene targets will be more difficult when analyzing products with complex mixtures of bacteria, making specificity analysis fundamental for validation of the method. Nowadays, several databases compiling full and partial rRNA sequences are available online and updated regularly, allowing fast and easy comparison of data from unknown organisms. These databases include the Ribosomal Database Project II Release 10 (update 14, released on August 31, 2009) that consists of 1074,075 aligned and annotated 16S rRNA sequences, along with seven online analysis tools (http://rdp. cme.msu.edu/). Amplification of the rRNA genes (e.g., PCR or RT–PCR) and the use of probes directed to these regions (e.g., FISH or microarrays) have been widely applied for the detection and/ or identification of Enterococcus as a genus or for particular species. Since E. faecalis is the most commonly encountered species, the majority of the primers and probes are specific for its identification. RNA Polymerase and Phenylalanyl-tRNA Synthase. In 2005, Naser106 and coworkers reported that all enterococcal species analyzed were clearly differentiated on the basis of their RNA polymerase (rpoA) and phenylalanyl-tRNA synthase (pheS) gene sequences. Strains of the same enterococcal species had at least 99% rpoA and 97% pheS sequence similarity, whereas different enterococcal species had a maximum of 97% rpoA and 86% pheS sequence similarity, pointing to the usefulness of this gene as a target for ­species-level

235

Enterococcus

TABLE 22.1 Probes for Detection and Identification of Enterococci Organism

Target

Eubacteria

16S rRNA 16S rRNA 16S rRNA 16S rRNA

EUB338- GCTGCCTCCCGTAGGAGT GTACAAGGCCCGGGAACGTATTCACC GACATAAGGGGCATGATGATTTGACGT Encl 31- CCCCTTCTGATGGGCAGG

FISH Microarrays Microarrays Microarrays

100 101 101 96

16S rRNAa 16S rRNA

Encl 45- GGGATAACACTTGCAAAC Encl 259- GAAGTCGCGAGGCTAAGC

Microarrays Microarrays

96 96

16S rRNA

ENC221- CACCGCGGGTCCATCCATCA

FISH

92

16S rRNA

ENF191- GAAAGCGCCTTTCACTCTTATGC

FISH

92

23S rRNA

TGGTTCTCTCCGAAATAGCTTTAGGGCTA

TaqMan PCR (qPCR)

88

23S rRNA

GPL813TQb- TGGTTCTCTCCGAAATAGCTTTAGGGCTA

TaqMan PCR (qPCR)

88

vanA enterococci

23S rRNA 23S rRNA vanA gene

Enc01aV- AGGTTAAGTGAATAAGGG ENC176- CAGTTCTCTGCGTCTACCTC vana3- CAACTAACGCGGCACTGTTTCCCAAT

Microarrays FISH TaqMan PCR (qPCR)

96 92 89

vanC enterococci

16S rRNA

EGAC183- CAACTTTCTTCCATGCGGAAAAT

FISH

92

E. casseliflavus

23S rRNA

Eca58- AGCTTGTCCGTACAGGTA

Microarrays

94

E. faecalis

16S rRNA

EFA1- TGATTTGAAAGGCGCTTTCGGGTGTCGCTGATGGATGGAC Microarrays

84

16S rRNA

EFA2- GAAGAACAAGGATGAGAGTAACTGTTCATCCCTTGACGG

Microarrays

84

16S rRNA

EFA3- GAAGTACAACGAGTTGCGAAGTCGCGAGGCTAAGCTAAT Microarrays

84

16S rRNA

CAATTGGAAAGAGGAGTGGCGGACG

TaqMan PCR (qPCR) 102

16S rRNA

TGCCGCATGGCATAAGAGTGAAAGGCGCTTTCGGGTGTC

Microarrays

103

16S rRNA

CAAGGACGTTAGTAACTGAACGTCCCCTGACGGTATCTAA

Microarrays

103

16S rRNA

GAAGTACAACGAGTCGTTAGACCGCGAGGTCATGCAAATC

Microarrays

103

16S rRNA

CCTCGCGGTCTAGCAGCTCGTTGTGCTT

Microarrays

101

16S rRNA

Enfl84- UGCACUCAAUUGGAAAGAGG

FISH

Gram-positive bacteria Enterococcus spp.

Sequence (5′–3′)

Method

Reference

91

16S rRNA

efs129- CCCTCTGATGGGTAGGTT

Microarrays

ITS region

Efs1- TATTTATTGATTAACCTTCT(T)

Hybridization

104

ITS region

Efs2- AAGAAGTGATCAAGACCCA(T)

Hybridization

104

23S rRNA

Efs18i- CGAAATGCTAACAACACC

Microarrays

modified by 93

23S rRNA

Efa54- CAAAAACAACTGGTACAG

Microarrays

96

16S rRNA

TGATTTGAAAGGCGCTTTCGGGTGTCGCTGATGGATGGAC

Microarrays

103

16S rRNA

GAAGAACAAGGATGAGAGTAACTGTTCATCCCTTGACGG

Microarrays

103

16S rRNA

GAAGTACAACGAGTTGCGAAGTCGCGAGGCTAAGCTAAT

Microarrays

103

ITS region

Efm1- TTTTATGAGACGATCGAT(T)

Hybridization

104

ITS region

Efm2- TCTTGATCTAACTTCTAT(T)

Hybridization

104

23S rRNA

Efm09i- GGATGTTACGATTGTGTG

Microarrays

modified by 93

23S rRNA

Efm09- CACACAATCGTAACATCC

Microarrays

96

23S rRNA

Efi58- TGACTCCTCTCCAGACTT

Microarrays

96

23S rRNA

ATCATACGATCAGCCGCAGTGAATA

RT–qPCR

76

23S rRNA

ENU140- TTCACACAATCGTAACATCCTA

FISH

92

23S rRNA

ENU1470- GACTCCTTCAGACTTACTGCTTGG

FISH

92

E. flavescens

23S rRNA

Efl58i- TTCTACCTATACGGACAA

Microarrays

E. gallinarum

ITS region

Egal- GAGTGGACAAGTTAAAGA(T)

Hybridization

23S rRNA

Ega09- CACAACTGTGTAACATCC

Microarrays

96

23S rRNA

EGA141- ATTCACAACTGTGTAACATCCTAT

FISH

92

23S rRNA

Eacdfg57- AGACATATCCATCAGTCT

Microarrays

96

ITS region

Ecasc- GAGTTGAAATGTTAAAAGAG(T)

Hybridization

23S rRNA

Ecafl09i- GGATGTTACGTCTGCGTG

Microarrays

E. faecium

E. flavescens/ E. gallinarum E. casseliflavus/ E. flavescens

96

96 104

104 96 continued

236

Molecular Detection of Human Bacterial Pathogens

TABLE 22.1 (Continued) Probes for Detection and Identification of Enterococci Organism

Target

Sequence (5′–3′)

Method

Reference

E. casseliflavus/ E. gallinarum/ E. flavescens E. faecium

16S rRNA

CGF- CCGTATAACACTATTTTCCGC

Hybridization

105

23S rRNA

Edfm57- CTGCTTGGACAGACATTT

Microarrays

E. faecium

16S rRNA

FMDUR- CATTCAGTTGGGCACTCTAGCAAGA

Hybridization

E. faecium

16S rRNA

Enfm93- CCGGAAAAAGAGGAGUGGC

FISH

91

E. gallinarum/ E. casseliflavus/ E. flavescens E. faecium

16S rRNA

Ecg191- GCGCCTTTCAACTTTCTT

Microarrays

96

16S rRNA

Enc93- GCCACTCCTCTTTTTCCG

Microarrays

96

E. casseliflavus/ E. flavescens

16S rRNA

Ecf459- GGGATGAACATTTTACTC

Microarrays

96

96 105

  Not E. faecalis.

a

TABLE 22.2 Broad-Range Control and Enterococcal Genus- and Species-Specific PCR Primers Primers (5′–3′)

Product size (bp)

Observations

Reference

Eubacteria 16S rRNA For- GGATTAGATACCCTGGTAGTCC

320

111 in 112

Rev- TCGTTGCGGGACTTAACCCAAC Sm785 For- GGATTAGATACCCTGGTAGTC

~1500

For- GCTGGATCACCTCCTTTC

676/578

Sm785 For + 422Rev

113 114

Uni331 For- TCCTACGGGAGGCAGCAGT

466

115

1554

84, 103

170

79

1522

86

996

116

Uni797 Rev- GGACTACCAGGGTATCTAATCCTGTT For- GAGAGTTTGATYCTGGCTCAG Rev- AAGGAGGTGATCCARCCGCA 1020 For- TTAAACTCAAAGGAATTGACGG 1190 Rev- CTCACGRCACGAGCTGACGAC Ef16 For- AGAGTTTGATCCTGGCTCA Ef16 Rev- GGTTACCTTGTTACGACTTC U1 For- CCAGCAGCCGCGGTAATACG U2 Rev- ATCGG(C/T)TACCTTGTTACGACTTC 23S rRNA 422 Rev- GGAGTATTTAGCCTT

~1500

Sm785 For + 422Rev

113

L189 Rev- GGTACTTABATGTTTCAGTTC

~1000

ENFE + L189

117

ECST784 For- AGAAATTCCAAACGAACTTG

92

118

900

76

ENC854 Rev- CAGTGCTCTACCTCCATCATT For- AGGAKGTTGGCTTAGAAGCAG Rev- CGCTACCTTAGGACCGTTATAGTTAC rrn 13B For- GTGAATACGTTCCCGGGCCT

~600

16S rRNA + 23S rRNA + ITS

104

6 Rev-GGGTTYCCCCRTTCRGAAAT Chaperonin 60 H279A For- GAIIIIGCIGGIGA(TC)GGIACIACIAC

600

109

H280A Rev- (TC)(TG)I(TC)(TG)ITCICC(AG)AAICCIGGIGC(TC)TT continued

237

Enterococcus

TABLE 22.2 (Continued) Broad-Range Control and Enterococcal Genus- and Species-Specific PCR Primers Primers (5′–3′) 590 For- GGNGACGGNACNACNACNGCAACNGT 590 Rev- TCNCCRAANCCNGGYGCNTTNACNGC

Product Size (bp)

Observations

Reference

589

110

803

83

Elongation factor EF-Tu (tuf) U1 For- AAYATGATIACIGGIGCIGCICARATGGA U2 Rev- AYRTTITCICCIGGCATIACCAT Lactic acid bacteria α subunit of ATP synthase (atpA) atpA-20 For- TAYRTYGGKGAYGGDATYGC

1102

107

494

106

431

106

815

106

atpA-27 Rev- CCRCGRTTHARYTTHGCYTG Phenylalanyl-tRNA synthase (pheS) pheS-21 For- CAYCCNGCHCGYGAYATGC pheS-22 Rev- CCWARVCCRAARGCAAARCC pheS-21 For- CAYCCNGCHCGYGAYATGC pheS-23 Rev- GGRTGRACCATVCCNGCHCC RNA polymerase a subunit (rpoA) rpoA-21 For- ATGATYGARTTTGAAAAACC rpoA-23 Rev- ACHGTRTTRATDCCDGCRCG Genus Enterococcus 16S rRNA 733

119

115

120

337

121

92

88

320 + 420

122,123 in 124

1609

110

Elongation factor EF-Tu (tuf) Ent1 For- TACTGACAAACCATTCATGATG Ent2 Rev- AACTTCGTCACCAACGCGAAC

112

83

Manganese-dependent superoxide dismutase (sodA) D1 For- CCITAYICITAYGAYGCIYTIGARCC D2 Rev- ARRTARTAIGCRTGYTCCCAIACRTC

480

82

E1 For- TCAACCGGGGAGGGT E2 Rev- ATTACTAGCGATTCCGG 16S rRNA Ec-ssu1 For- GGATAACACTTGGAAACAGG Ec-ssu1 Rev- TCCTTGTTCTTCTCTAACAA 16S rRNA For- ATCAGAGGGGGATAACACTT Rev- ACTCTCATCCTTGTTCTTCTC 23S rRNA ECST784 For - AGAAATTCCAAACGAACTTG ENC854 Rev- CAGTGCTCTACCTCCATCATT ITS region For- CAAGGCATCCACCGT Rev- GAAGTCGTAACAAGG Chaperonin 60 GroES EntGroES For- TTAAAACCATTAGGCGATCG EntGroES Rev- CCCATNCCCATNGANGGRTCCAT

E. faecalis 16S rRNA Efs130 For- AACCTACCCATCAGAGGG Efs490 Rev- GACGTTCAGTTACTAACG For- CGCTTCTTTCCTCCCGAGT Rev- GCCATGCGGCATAAACTG

360 143

125 126 102 continued

238

Molecular Detection of Human Bacterial Pathogens

TABLE 22.2 (Continued) Broad-Range Control and Enterococcal Genus- and Species-Specific PCR Primers Primers (5′–3′)

Product Size (bp)

For- TACTGACAAACCATTCATGATG

Observations

Reference

112

  127

138

79

Rev- AACTTCGTCACCAACGCGAAC 72 For- CCGAGTGCTTGCACTCAATTGG 210 Rev- CTCTTATGCCATGCGGCATAAAC ENFE For- GTCGCTAGACCGCGAGGTCATGA

~1000

ENFE + L189

128

Chaperonin 60 GroESL 185

110

64

76

650

86

941

80

475

129

803

86

360

108

444

77

518

130

EE1 For- TGTGGTATCGGAGCTGCAG

430

129

EE2 Rev- ATAGTTTAGCTGGTAAC For- TGTGGTATCGGAGCTGCAG Rev- GTCGATTCTCGCTAATCC

513

131

810

132

941

129

EfGroES For- GGAATTGTTCTTGCATCCGT EfGroES Rev- ACAATTAAGTATTCTACGCC For- TGTGGCAACAGGGATCAAGA Rev- TTCAGCGATTTGACGGATTG Efes For- GTGTTAAAACCATTAGGCGAT Efes Rev- AAGCCTTCACGAACAATGG d-Alanine/d-alanine

ligase (ddl)

For- ATCAAGTACAGTTAGTCTTTATTAG Rev- ACGATTCAAAGCTAACTGAATCAGT DD13 For- CACCTGAAGAAACAGGC DD13 Rev- ATGGCTACTTCAATTTCACG Iron sulfur–binding protein Efis For- ATGCCGACATTGAAAGAAAAAATT Efis Rev- TCAATCTTTGGTTCCATCTCT Manganese-dependent superoxide dismutase (sodA) FL1 For- ACTTATGTGACTAACTTAACC FL2 Rev- TAATGGTGAATCTTGGTTTGG Penicillin-binding protein (pbp5) For- CATGCGCAATTAATCGG Rev- CATAGCCTGTCGCAAAAC Ef0027- unknown function For- GCCACTATTTCTCGGACAGC Rev- GTCGTCCCTTTGGCAAATAA vane

vanG For- CGGTTGTGCCGTACTTGGC Rev- GGGTAAAGCCATAGTCTGGGGC EG1 For- CGGCATCCGCTGTTTTTGA EG2 Rev- GAACGATAGACCAATGCCTT

E. faecalis and E. faecium vanA vana3 For- CTGTGAGGTCGGTTGTGCG vana3 Rev- TTTGGTCCACCTCGCCA

64

89

vanB For- GTGACAAACCGGAGGCGAGGA Rev- CCGCCATCCTCCTGCAAAAAA

433

133

For- CATCGCCGTCCCCGAATTTCAAA

298

81

Rev- GATGCGGAAGATACCGTGGCT continued

239

Enterococcus

TABLE 22.2 (Continued) Broad-Range Control and Enterococcal Genus- and Species-Specific PCR Primers Primers (5′–3′)

Product Size (bp)

EB3 For- ACGGAATGGGAAGCCGA

Observations

647

Reference 129

EB4 Rev- TGCACCCGATTTCGTTC For- CATCGCCGTCCCCGAATTTCAAA

297

134

500

129

630

135

461

136

628

137

Rev- GATGCGGAAGATACCGTGGCT vanD ED1 For- TGTGGGATGCGATATTCAA ED2 Rev- TGCAGCCAAGTATCCGGTAA For- GARGATGGITSCATMCARGGW Rev- MGTRAAICCIGGCAKRGTRTT For-TAAGGCGCTTGCATATACCG Rev- TGCAGCCAAGTATCCGGTAA vanD-U1 TATTGGAATCACAAAATCCGG vanD-U2 CGGCTGTGCTTCCTGATG E. faecium 23S rRNA Ef Rev- CACACAATCGTAACATCCTA d-Alanine/d-alanine

676/578

95

550

80

1091

129

215

108

658

138

ligase (ddl)

For- GCAAGGCTTCTTAGAGA Rev- CATCGTGTAAGCTAACTTC FAC1 For- GAGTAAATCACTGAACGA FAC2 Rev- CGCTGATGGTATCGATTCAT Manganese-dependent superoxide dismutase (sodA) FM1 For- GAAAAAACAATAGAAGAATTAT FM2 Rev- TGCTTTTTTGAATTCTTCTTTA Unknown region For- TTGAGGCAGACCAGATTGACG Rev- TATGACAGCGACTCCGATTCC

Other enterococcal species Manganese-dependent superoxide dismutase (sodA) 288

E. casseliflavus

108

FV1 For- GAATTAGGTGAAAAAAAAGTT

284

E. flavescens

108

FV2 Rev- GCTAGTTTACCGTCTTTAACG GA1 For- TTACTTGCTGATTTTGATTCG GA2 Rev- TGAATTCTTCTTTGAAATCAG

173

E. gallinarum

108

1030

Several enterococcal species

133

377

Several enterococcal species

81

783

Several enterococcal species

134

822

E. gallinarum and E. casseliflavus/E. flavescens

80

438

E. gallinarum and E. casseliflavus/E. flavescens

81

CA1 For- TCCTGAATTAGGTGAAAAAAC CA2 Rev- GCTAGTTTACCGTCTTTAACG

vanA For- CATGAATAGAATAAAAGTTGCAATA Rev- CCCCTTTAACGCTAATACGATCAA For- TCTGCAATAGAGATAGCCGC Rev- GGAGTAGCTATCCCAGCATT For- GCTATTCAG CTGTACTC Rev- CAGCGGCCATCATACGG vanC1 For- GGTATCAAGGAAACCTC Rev- CTTCCGCCATCATAGCT For- GACCCGCTGAAATATGAAG Rev- CGGCTTGATAAAGATCGGG continued

240

Molecular Detection of Human Bacterial Pathogens

TABLE 22.2 (Continued) Broad-Range Control and Enterococcal Genus- and Species-Specific PCR Primers Primers (5′–3′) For- GGTATCAAGGAAACCTC

Product Size (bp)

Observations

Reference

822

E. gallinarum and E. casseliflavus/E. flavescens

134

815/827

E. gallinarum and E. casseliflavus/E. flavescens

129

430

E. gallinarum and E. casseliflavus/E. flavescens

81

484

E. gallinarum and E. casseliflavus/E. flavescens

139

439

E. gallinarum and E. casseliflavus/E. flavescens

134

Rev- CTTCCGCCATCATAGCT vanC1/C2 EC5 For- ATGGATTGGTAYTKGTAT EC8 Rev- TAGCGGGAGTGMCYMGTAA vanC2 For- CTCCTACGATTCTCTTG Rev- CGAGCAAGACCTTTAAG vanC2/C3 For-CGGGGAAGATGGCAGTAT Rev- CGCAGGGACGGTGATTTT For- CTCCTACGATTCTCTTG Rev- CGAGCAAGACCTTTAAG

molecular identification. However, no species-specific primers have yet been developed or applied to this genus. Elongation Factor EF-Tu. The tuf gene, encoding elongation factor EF-Tu, is involved in peptide chain formation and is an essential constituent of the bacterial genome83,141 making this region a good target for diagnostic purposes. Ke and coworker83 reported in 1999 a PCR-based assay that targets the tuf gene and detects most enterococcal species with excellent sensitivity and acceptable specificity. d-Alanine:d-Alanine Ligase. A PCR assay developed by Dutka-Malen and coworkers80 enabling the identification at the species level (species E. faecium and E. faecalis) and targeting the d-alanine:d-alanine ligase (ddl) gene was applied to characterize clinically relevant enterococci. Primers for speciesspecific PCR of E. durans and E. hirae targeting the ddl gene were also developed and validated by Knijff and coworkers.142 Penicillin-Binding Proteins. Penicillin-binding proteins (PBPs) are the enzymes involved in terminal stages of peptidoglycan synthesis, and PBP1 and PBP5 are the prevalent PBPs. Thus, these proteins are potential targets for bacterial cell detection. The gene coding for PBP5 (pbp5) has previously been used for the development of an E. faecalis speciesspecific probe143 and was subsequently used in competitive PCR for quantification of nonculturable E. faecalis cells.77 Muramidase. In 2006, Arias and coworkers87 searched the GenBank for sequences to be applied for the identification of E. hirae. During this process, an E. durans muramidase gene (mur) with 82% homology to E. hirae was detected; primers were developed for both genes and used for multiplex PCR. This method appears to provide a rapid and accurate identification of these two very similar enterococcal species. ATP Synthase. atpA codes for the alpha subunit of the bacterial ATP synthase, which functions in ATP synthesis

coupled to proton transport.144 The aim of the study developed by Naser and coworkers107 was to analyze the usefulness of atpA gene sequences for the reliable identification of Enterococcus species. All species were differentiated, with a maximum of 92% similarity. The intraspecies atpA sequence similarities for all species varied from 98.6% to 100%, except for E. faecium strains that showed a lower atpA sequence similarity of 96.3%. This study clearly showed that atpA provides an alternative tool for the phylogenetic study and identification of enterococci, but no genus- or speciesspecific primers have been described to demonstrate this potential. Manganese-Dependent Superoxide Dismutase. In 2000, Poyart and coworkers82 sequenced the manganesedependent superoxide dismutase gene (sodA) of 19 species of the genus Enterococcus. Since variations in their sequences appeared to be greater between species and lower within species, this gene was used to develop species-specific primers108 that can be applied to identify 23 validly published enterococcal species. Heat-Shock Proteins. The groESL genes (also known as chaperonin 60 genes, cpn10/60 or hsp10/60), which encode 10-kDa (GroES) and 60-kDa (GroEL) heat-shock proteins, are ubiquitous and evolutionarily highly conserved among bacteria.145 Goh and coworkers109 demonstrated that reverse checkerboard hybridization methodology, based on an approximately 600-bp region of the chaperonin 60 gene, can be an efficient method for the identification of Enterococcus species. Teng and coworkers110 determined the groESL gene’s full-length sequence of E. faecalis and used this information to develop primers specific for this species. Antimicrobial Resistance. Over the years, several authors used multiplex PCR to both detect the presence of vancomycin

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resistance determinants and identify enterococci to the species level.80,112,129,139,146 Glycopeptide resistance has been detected in several species of enterococci, with seven glycopeptide resistance genotypes already described. Five of these genotypes (vanA, vanB, vanD, vanE, and vanG) are acquired mechanisms and the other two (vanC1 and vanC2/C3) are intrinsic properties. vanA and vanB are the most common resistance genotypes. The vanA genes encode proteins that confer high-level resistance to vancomycin and teicoplanin147 and have been found in several enterococcal species.148 The vanB genes confer resistance to various concentrations of vancomycin but not teicoplanin149; vanB-type glycopeptide resistance has been described for E. faecalis and E. faecium. The vanD phenotype is characterized by resistance to moderate levels of vancomycin and to low levels of teicoplanin;136,150 vanD determinants are constitutively expressed in E. faecalis but inducible in E. faecium.136 The vanE gene is induced by low levels of vancomycin131 and has only been detected in one strain of E. faecalis. The vanC1 and vanC2/C3 genes are specific to the motile VRE species E. gallinarum and E. casseliflavus/E. flavescens, respectively.80 Recently, a new vancomycin resistance locus, vanG, has been detected in four E. faecalis clinical isolates with a moderated level of resistance to vancomycin and full susceptibility to teicoplanin.132

22.2  METHODS 22.2.1  Sample Preparation It is often difficult to establish whether an Enterococcus strain is contributing to the infection or whether it is colonizing a human or an animal. Bacteremia as major cause of nosocomial infection represents an important source of morbidity and mortality. Gram-positive microorganisms, especially Staphylococcus and E. faecalis, account for the majority of episodes of bacteremia in critically ill patients in intensive care units (ICU). The rapid and reliable detection and subsequent identification of microorganisms, namely, Enterococcus, from blood is critical so that appropriate antimicrobial therapy can be provided. The same principle has to apply to the rapid and reliable detection of microbial samples from stool samples, oral samples, urine, pus, or exudates. The appropriate collection and transport of specimens are critical to ensure that proper diagnostic information is given by the microbiology laboratory.151,152 Also the initial processing of clinical specimens, which is a multifaceted endeavor as one has to consider specimen type and its anatomical origin, is a critical step. Description of the pretreatment and treatment of the specimens received in laboratory for aerobic microbiology in which Enterococcus is included have been gathered by York in 2007,153 but generally they start by microscopical observation, Gram stain and isolation from the clinical samples in liquid and solid media, followed by identification procedures. Selective agents that are commonly used in selective media to isolate enterococci include: sodium azide, thallous acetate, kanamycin, and gentamicin. Other limiting ingredients or factors are crystal violet, Tween

80, carbonate, and bile salts. Special growth conditions may be also used, like low pH (e.g., 6.0 or 6.2) or elevated incubation temperature (42°C or 45°C). Nutritional requirements vary among the members of the genus, but it usually an advantage to use peptones, beef or yeast extract. Considering the mentioned factors it is easy to understand why there are so many varieties of selective media and discrepant results in the literature. ABA (esculin bile azide agar), KAA (kanamycin aesculin azide agar), CATC (citrate azide tween carbonate agar), ME (m-Enterococcus agar), TITG (thallous acetate tetrazolium glucose agar), and KA (crystal violet azide agar) are commonly used to cultivate enterocci.154–156 Laboratories vary on the media they use depending on the type of sample; for instance, some use KAA agar to isolate enterococci from feces and cysteine-lactose-electrolyte-deficient (CLED) agar can be used in bacteriological studies of urine.157 The detection and identification of vancomycin-resistant enterococci (VRE) led to the development of special media and procedures to detect and identify them. Some authors demonstrated that VRE grew on Mueller-Hinton agar with 6–12 µg/mL of vancomycin, although 6 µg/mL of vancomycin seems to be the optimum concentration.12,158 Chromogenic media are now also widely used namely for cell counts and identification of enterococci from urine or stool samples, either if they are vancomycin-resistant or not. Media like BBL CHROMagar (BD Diagnostics), Chromogenic VRE (AES Laboratoire), chromID VRE (bioMérieux), VRE agar (Oxoid) or even bile esculin agar supplemented with 6 µg/mL of vancomycin (BEV), and bile esculin azide vancomycin (BEAV) agar have been used.159–162 After isolation of potential Enterococcus strains from clinical samples, using any of the media already mentioned, strains can be grown in trypticase-soy-5% sheep blood agar, brain heart infusion-5% sheep blood agar, or any blood agar base containing 5% animal blood, as well as brain heart infusion (BHI), MRS agar or broth (DeMan, Rogosa, Sharpe) or M17agar or broth to be further characterized or identified by PCR. All enterococci grow at 35ºC–37ºC and do not require an atmosphere containing increased levels of carbon dioxide.

22.2.2  Detection Procedures 22.2.2.1  Semiautomatic and Automatic Identification The blood culture is the gold standard for diagnosing bacteria, but it may take more than two days for results to be available. Semiautomatic or automatic methods to test blood culture or other body fluids and to identify enterococci directly from samples, like BACTEC Peds PLUS/F and BACTEC 9240 that use spectrophotometry to monitor the CO2 being produced during growth, are now commonly used, and the results have been compared to other systems of identification.163,164 Other systems used for the identification of clinically isolated enterococci are kits like API 20 Strep and API Rapid ID 32 STREP (bioMérieux) and BBL Crystal gram-positive and BBL Crystal Rapid gram-positive (BD Diagnostics), that were already compared in terms of

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specificity.165 There are also systems that identify and at the same time test the antibiotic susceptibility of the strains like the Vitek 2 and Vitek Compact Systems (bioMerieux). Also very used are the identification systems Bact/ALERT 3D (bioMérieux), BACTEC 9000 (BD Diagnostics), Versa Trek (Trek), or Phoenix 100 (Becton Dickson).165–168 22.2.2.2  M  ultiplex PCR for the Detection of Enterococcus Species Although most of the routine identification of pathogens in clinical samples start with bacterial isolation, the advent of molecular approaches opens the road for direct application of such methods without culturing. In this context, the experimental procedure hereby described may be applied using DNA extracted both from isolated strains and clinical samples. A list of kits that can be used to extract DNA, directly from biological samples or from isolated colonies, has already been compiled.140 After DNA extraction, it is proposed that the detection/ identification of Enterococcus can follow the steps shown in Clinical sample

DNA extraction

Primers

E1 + E2 FL1 + FL2 FM1 + FM2

Target species

Identification by genus- and species-specific multiplex PCR

E. faecalis 360 bp

E. casseliflavus 288 bp

E. faecium 215 bp

E. gallinarum 173 bp

E1 + E2 CA1 + CA2 GA1 + GA2

Screening of putative virulence factors by multiplex PCR

A1 + A2 B1 + B2 C1 + C2 D1 + D2

CYT I + CYT IIb ESP 14F + ESP 12R ASA 11 + ASA 12 GEL 11 + GEL 12 HYL n1 + HYL n2

Target genes

Primers

Screening of antimicrobial resistance by multiplex PCR

vanA vanB vanC1 vanC2/C3

cytolysin -cylAenterococcal surface protein -espaggregation substance -asalgelatinase -gelEhialuronidase -hyl-

FIGURE 22.1  Flowchart of the multiplex-PCR-based procedures for the detection and/or identification of enterococci at genus and species level, as well as for the assessment of antimicrobial and virulence determinants.

the flow chart of Figure 22.1. The procedure is a genus- and species-specific multiplex PCR, adapted from the 23-species detection methodology of Jackson and coworkers,108 enabling the detection of E. faecalis, E. faecium, E. casseliflavus, and E. gallinarum. Because this approach combines the genus-specific primers (E1/E2) described by Deasy and coworkers119 with primers for the superoxide dismutase (sod) gene108 specific for E. faecalis (FL1/FL2), E. faecium (FM1/FM2), E. casseliflavus (CA1/ CA2), and E. gallinarum (GA1/GA), it provides a rapid and reliable method for the detection and/or identification of the most important human pathogenic enterococcal species. Amplification procedure consists of the preparation of a PCR master mix (60 µL) containing 3 mM MgCl2, 0.2 mM dNTP mix, 3.5 U of high fidelity DNA polymerase, and 3.75  μL (16 μM) of genus-specific primer set (E1/E2). The master mix should be equally divided in three separated vials in order to use the primer combinations depicted in Figure 22.1 (for sequence see Table 22.2) and have a positive control of the reaction. To vial number 1 add 2.5 µL (16 µM) of primers FL1/FL2 and 1.25 μL (16 μM) of primers FM1/FM2; to vial number 2 add 2.5 µL (16 µM) of primers GA1/GA2 and 1.25 μL (16 μM) of primers CA1/CA2; to vial number 3 (positive control) add DNA from a nonpathogenic strain of Enterococcus. PCR reactions are performed in a final volume of 22.5 μL after the addition to each vial of 2.5 μL of test DNA, extracted either from clinical sample or isolated strain. The PCR amplification program includes an initial denaturation at 95°C for 4 min; 30 cycles of 95°C for 30 s, 55°C for 1 min, and 72°C for 1 min; a final extension step at 72°C for 7 min. After PCR amplification an aliquot of 10 μL of product is electrophoresed on a 2% 1× Tris-acetate-EDTA agarose gel containing 2 μg/mL ethidium bromide. A DNA molecular size marker (preferably with a 10 bp resolution) should be used as standard. Following electrophoresis, the agarose gels are visualized and photographed under UV light. Since the presence of enterococcal isolates carrying virulence and antimicrobial resistance determinants turns the pathogenicity potential of these bacteria higher, we suggest an integrative approach leading to species identification and assessment for the presence of vancomycin resistance and virulence determinants (Figure 22.1). Although this approach is not essential for treating enterococcal infections, it may contribute to the implementation of more adequate antimicrobial treatment and/or to the implementation of measures aimed at the elimination of more virulent enterococci from hospital settings. Screening for the presence of antimicrobial resistance and virulence determinants can also be performed by multiplex PCR, using the methodologies described by DutkaMalen et al.80 and Vankerckhoven et al.,169 respectively. Using the molecular size marker included in the agarose gel, the size of the amplicons can be estimated and compared with the expected ones (Table 22.2 and Figure 22.1). If the obtained fragments show the expected molecular size for the genus and species primer set, the presence of the target species is detected in the sample (or the isolate is identified at species level if working with pure cultures isolated from clinical

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Enterococcus

samples). When only the genus-specific amplicon is visualized, the presence of another enterococcal species is inferred. If no amplicons are observed, with the exception of the positive control reaction tube (which demonstrates the absence of PCR inhibitors), this means that either no enterococci are present in the analyzed sample at detectable levels or the existing strains show sequence variations in sod gene that prevent primer annealing (the false-negative problem can occur despite the applied methodology and has to be taken in account).

22.3  C  ONCLUSIONS AND FUTURE PERSPECTIVES Enterococcus are generally not as virulent as other grampositive bacteria but can occur as a polymicrobial infection in debilitated hosts.170 Resistance of enteroccci to multiple antimicrobial agents, including vancomycin, can occur in health care centers. Considering that the most problematic enteroccci are VREs, the infection control has to pass through careful analysis on VRE spread. Since colonized patients are the primary reservoir of VRE, a patient-to-patient transmission occurs in the first place. As hospital staff and environmental contamination, through medical devices or surroundings of colonized patients, also plays a critical role in spread, all these aspects have to be taken into consideration.170 The Hospital Infection Control Practices Advisory Committee (HIPAC) has published in 1995 “Recommendations for Preventing the Spread of Vancomycin Resistance” (Centers for Disease Control and Prevention. Recommendations for Preventing the Spread of Vancomycin Resistance).171 These measures include the appropriate use of vancomycin, the establishment of education programs, the clear understanding of the role of microbiology laboratory, the prevention and control of nosocomial transmission of VRE, and the isolation precautions to prevent patient-to-patient transmission. More recently, research on antimicrobial stewardship, meaning the optimal selection dose and duration of an antimicrobial that results in best clinical outcome for the treatment and prevention of infection, with minimum toxicity to the patient and minimal impact on subsequent resistance and development, has produced more and more data.172–174 Health care facilities should then adopt antimicrobial stewardship programs (ASPs) and infection control programs (ICPs) to monitor antimicrobial use while simultaneously optimizing treatment, outcome, and cost. The IDSA/SHEA guidelines for developing an institutional program to enhance antimicrobial stewardship can be used as a starting point to control the threat.175,176 All these evaluations have to take on board, not only healthy people, who seem to be unaffected by enterococcal strains, but also the elderly population, infants, and immunocompromised people. The microbiological risk assessment of emerging pathogens has to be performed, and global data indicate that the epidemiology of human diseases is changing because of changes in lifestyles and improved surveillance and recovery/detection methods.177,178 Pathogen control, and especially VRE, depends on the laboratory’s ability to detect,

identify, and recognize those agents; therefore, in the case of Enterococcus, a detailed combination of a more specific genus and species method is proposed here based on the work of Jackson and coworkers.108 PCR methods in general, and real-time PCR methods in particular, are increasingly used for the detection and quantification of pathogens in clinical samples. Nonetheless, the new species that are constantly appearing in the literature, calling for new and/or reassessed methods of identification and quantification as well as for the need to always be alert to new antibiotic resistances, are a constant challenge and constitute new routes of needed research in the area of work with Enterococcus.

ACKNOWLEDGMENTS The authors acknowledge the partial funding from different projects on Enterococcus or food quality and safety that have financed their research on the subject, namely, Fundação para a Ciência e Tecnologia through Program PRAXIS XXI, project 2/2.1/BIO/1121/95 and Program SAPIENS, project POCTI/AGR/36165/99 and EU, through Specific Supported Action project BIOSAFE SSPS-CT-2006–022725. T. Semedo-Lemsaddek thanks the financial support of the FCT Pos-Doctoral fellowship SFRH/BPD/20892/2004.

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23 Granulicatella Sheng Kai Tung and Tsung Chain Chang CONTENTS 23.1 Introduction...................................................................................................................................................................... 249 23.1.1 Classification, Morphology, and Biology............................................................................................................. 249 23.1.2 Clinical Features................................................................................................................................................... 249 23.1.3 Diagnosis.............................................................................................................................................................. 250 23.1.3.1 Conventional Techniques....................................................................................................................... 250 23.1.3.2 Molecular Techniques............................................................................................................................ 251 23.2 Methods............................................................................................................................................................................ 251 23.2.1 Sample Preparation............................................................................................................................................... 251 23.2.2 Detection Procedures............................................................................................................................................ 251 23.2.2.1 16S rRNA Gene Sequencing.................................................................................................................. 251 23.2.2.2 Sequencing of the Ribosomal 16S-23S Intergenic Spacer (ITS) Region............................................... 252 23.2.2.3 Partial rpoB Gene Sequencing.............................................................................................................. 252 23.2.2.4 Array-Based Method.............................................................................................................................. 252 23.3 Conclusion and Future Perspectives................................................................................................................................. 253 References.................................................................................................................................................................................. 254

23.1  INTRODUCTION 23.1.1  Classification, Morphology, and Biology Bacteria in the genus Granulicatella are gram-positive, catalase-negative, facultative anaerobic cocci. Species in the genus can cause opportunistic infections in immunocompromised patients and are infrequently isolated from clinical laboratories.1 The genera Abiotrophia and Granulicatella have been proposed to accommodate microorganisms previously known as nutritionally variant streptococci or satelliting streptococci.1–3 By chromosomal DNA–DNA hybridizations, Bouvet et al.4 demonstrated that nutritionally variant streptococci were really members of two novel streptococcal species, Streptoccoccus defectivus and Streptoccoccus adjacens. Based on the 16S rRNA gene sequence, Kawamura et al.3 transferred Streptoccoccus defectivus and Streptoccoccus adjacens, respectfully, to two new species, Abiotrophia defectiva and Abiotrophia adiacens. A third species, Abiotrophia elegans, a possible pathogen in patients with endocarditis, was described in 1998 by Roggenkamp et al.5 In 2000, Collins and Lawson2 proposed a new genus, Granulicatella, with Granulicatella adiacens and Granulicatella elegans encompassing strains formerly recognized as Abiotrophia adiacens and Abiotrophia elegans, respectfully. Abiotrophia defectiva remains as the sole species in Abiotrophia.2 A third species in Granulicatella is Granulicatella balaenopterae; however, there was no report of human infection caused by this microorganism.

Bacteria in the genus Granulicatella are nonmotile cocci that occur as single cells, in pairs, or in short chains. Cell morphology depends on culture conditions. Elongated or swollen cells may be observed, especially when grown under less optimal nutritional conditions.6 The G + C content of Granulicatella DNA is approximately 36–37.5 mol%. Members of Granulicatella are normal flora of the oral cavity or upper respiratory tract1 and are non-betahemolytic on blood-based solid media. Granulicatella exhibit satellitism around the colonies of other bacteria, such as Staphylococcus epidermidis and other streptococci.7 Granulicatella species usually have low virulence and are normally pathogenic in immunocompromised hosts.1 Predisposing factors that lead to Granulicatella infections include previously damaged tissues (e.g., heart valves), invasive procedures, prolonged hospitalization, and the presence of foreign bodies. Granulicatella can be isolated from a variety of clinical samples, including blood, urine, cerebrospinal fluid, wound tissues, and others specimens.1 Infections caused by Granulicatella might be due to some microorganisms entering the bloodstream and disseminating to other tissues.8,9

23.1.2  Clinical Features Granulicatella species have been reported as etiologic agents of bacteremia, infectious endocarditis, brain abscess, postpartum or postabortal sepsis, pancreatic abscess, wound 249

250

infection, vertebral osteomyelitis, conjunctivitis, cirrhosis, endophthalmitis, infectious crystalline keratopathy, septic arthritis, otitis media, central nervous system infections, breast implant–associated infection, and peritoneal dialysis– related peritonitis.5,7,10–17 Using a molecular technique, the proportion of Granulicatella elegans, Granulicatella adiacens, and Abiotrophia defectiva was found to be 1:11:1 in the human mouth.18 Exopolysaccharides that can adhere to human tissues are produced by Granulicatella.7 All members of Granulicatella are susceptible to vancomycin. However, there are still no standardized methods and interpretation criteria for antibiotic susceptibility testing results in the documents of CLSI (Clinical and Laboratory Standards Institute). Most investigators have employed streptococcal (other than Streptococcus pneumoniae) interpretive criteria for determining the results of susceptibility.1 Since Granulicatella species are fastidious, Mueller-Hinton medium supplemented with blood and pyridoxal hydrochloride (0.001%) is used for susceptibility testing, and a CO2-enriched atmosphere1 is required for incubation. Tuohy et al.19 examined a collection of 21 Granulicatella adiacens isolates for their susceptibilities to a variety of antibiotics. They found that all strains were susceptible to clindamycin, rifampin, levofloxacin, ofloxacin, and quinupristin-dalfopristin. The percentage of isolates that were susceptible to other agents was as follows: penicillin, 55%; amoxicillin, 81%; ceftriazone, 63%; and meropenem, 96%. However, Zheng et al.20 observed high rates of beta-lactam and macrolide resistance in a collection of pediatric Granulicatella isolates, and hence they proposed that antimicrobial susceptibility testing should be systematically done to achieve appropriate antimicrobial therapy. In severely ill patients or those with a suboptimal response to initial therapy with beta-lactam antibiotics, treatment with vancomycin should be considered.21 The combination therapy with vancomycin and an aminoglycoside should be initiated empirically in patients with central nervous system infections.13

23.1.3  Diagnosis 23.1.3.1  Conventional Techniques Phenotypic characteristics that can be used as a guide to identify Granulicatella and Abiotrophia species are listed in Table 23.1. Members of Granulicatella are likely to be isolated on rich and nonselective media. They can grow on chocolate agar, in thioglycolate broth, and on brucella agar with 5% horse blood, but not on Typticase soy agar with 5% sheep blood.1 Cell growth requires thiol compounds such as l-cysteine or the active form of vitamin B6, pyridoxal phosphate, in media. Growth does not occur at 10°C or 45°C. Granulicatella in clinical samples may be overlooked when media without nutritional supplements are used. Gram stain morphology, although prone to be subjective in interpretation, is a major decision point in the identification protocols. Gram stain morphology of Granulicatella spp. resembles that of streptococci, with cells being cocci or coccobacilli in pairs and chains. Cells grown in broth, such as thioglycollate, should be used for determination of cellular morphology.

Molecular Detection of Human Bacterial Pathogens

Besides the characteristics listed in Table 23.1, additional tests are recommended to make a final identification, especially for isolates recovered from important clinical specimens.1 Additional tests useful for Granulicatella spp. differentiation can be found in several studies.4,6,22 Commercial identification kits, such as API 20 Strep, rapid ID32 STREP, and the VITEK 2 system (all from BioMérieux, Marcy l’Etoile, France), could be used for identification of Granulicatella spp. However, Granulicatella elegans cannot be identified by both API 20 Strep and rapid ID32 STREP kits since the species is not included in the databases of the two kits. Granulicatella adiacens and Granulicatella elegans are phenotypically similar; several tests (hydrolysis of arginine and hippurate, and production of β-glucuronidase) can be used to differentiate the two species (Table 23.1). Most clinical Granulicatella adiacens isolates could be identified by the API 20 STREP system with > 80%–90% confidence, whereas both the Vitek System (GPI) and ID32 STREP identified only a small portion of Granulicatella adiacens isolates.23 It should be noted that the growth of Granulicatella elegans is only supported by l-cysteine hydrochloride and not by pyridoxal phosphate when the organism is grown in ToddHewitt or casein-soy peptone broth.5 The media of blood culture systems supplemented only with pyridoxal phosphate TABLE 23.1 Phenotypic Characteristics of Abiotrophia defectiva and Granulicatella Speciesa Characteristics A. defectiva G. adiacens G. balaenopterae G. elegans Production of acid from Lactose Maltose Pullulan Sucrose Tagatose Trehalose

+ + + + − +

− + − + + −

− + + − − +

− − − + − −

Hydrolysis of Arginine Hippurate

− −

− −

+ −

+ +

Production of a-Galactosidase

+

−

−

−

β-Galactosidase

+

−

−

−

β-Glucuronidase

−

+

−

−

N-Acetyl-βglucosaminidase Pyrrolidonyl arylamidase Leucine aminopeptidase Satelliting behavior

−

−

+

−

+

+

ND

+

+

+

ND

+

+

+

ND

+

Symbols: −, negative reactions ≤15%; +, positive reactions ≥85%; ND, not determined. a  Data compiled from References 1, 2, 5, 6, 44, and 45.

251

Granulicatella

may fail to support the growth of Granulicatella elegans, and thus, these systems might not be able to detect this microorganism as a possible pathogen. 23.1.3.2  Molecular Techniques Molecular methods developed for identification of members of Granulicatella are limited. Sequence analysis of the 16S rRNA gene has been widely used for bacterial identification.24,25 The technique is becoming popular for identifying biochemically unidentified bacteria or for providing reference identifications for unusual strains.24 Sequencing of the 16S rRNA gene can be successfully used to identify Granulicatella species. Multiple sequences of the 16S rRNA genes for Granulicatella adiacens and Granulicatella elegans are available in the nucleotide database of National Center for Biotechnology Information, USA. The 16S rRNA gene sequences of Granulicatella adiacens and Abiotrophia defectiva formed a distinct cluster; this cluster is not closely related to other viridans streptococcal species and Abiotrophia.3,5 Woo et al.23 found that 16S rRNA gene sequencing is the technique of choice for identifying Granulicatella adiacens and proposed that early surgical intervention should be considered when endocarditis caused by this microorganism is diagnosed. The ribosomal 16S-23S intergenic spacer (ITS) region has been suggested as a good candidate for bacterial identification and strain typing.26–29 Sequences of the ITS region in bacteria have low intraspecies variation and high interspecies divergence.26,27,29 The feasibility of using ITS sequence to identify Granulicatella adiacens and Granulicatella elegans was established by Tung et al.30 It should be noted that in the genome of Granulicatella spp. possesses multiple ITS fragments having different lengths and sequences, therefore the PCR products amplified from the ITS region of an isolate cannot be directly sequenced. Before sequencing, the amplicons should be eluted from agarose gel or cloned into a plasmid and reamplified. Once the ITS sequence is determined, species identification can be done by searching databases using the BLAST (http://www.ncbi.nlm.nih.gov/ blast/) sequence analysis tool. The rpoB gene, encoding the highly conserved subunit of the bacterial RNA polymerase, has been demonstrated to be a suitable target for identification of a wide range of bacteria.31 The gene was shown to be more discriminative than the 16S rRNA gene for identifying enteric bacteria.31 Partial rpoB gene sequence (740 bp) had been applied to identify streptococci and Granulicatella adiacens.32 In recent years, DNA array technology has been successfully used to identify microorganisms that are difficult to be determined to the species level; these bacteria include viridans streptococci,33 nonfermenting gram-negative bac­ teria,34 Legionella,35 Mycobacterium36,37 and foodborne bacterial pathogens.38 The method consists of PCR amplification of a specific gene region, followed by hybridization of the PCR product labeled with a fluorescent dye (or with a molecular technique that could be recognized by antibody) to a panel of oligonucleotide probes immobilized on a solid

support such as glass slide or nylon membrane. The test sensitivity and specificity of the arrays are largely determined by the sequences of the oligonucleotide probes. The whole procedure of array hybridization takes between 8 and 24 h, starting from isolated colonies or clinical specimens. The hybridization patterns could be read by a fluorescence scanner if a fluorescent label is used or by the naked eye if colorimetric reaction is used. By using the oligonucleotide array platform, five oligonucleotide probes designed from the ITS regions were successfully used to identify Granulicatella adiacens, Granulicatella elegans, and Granulicatella balaenopterae.39 The array method used a protocol encompassing DNA extraction, PCR amplification of the ITS regions, and hybridization of the PCR products to speciesspecific probes on the array.

23.2  METHODS 23.2.1  Sample Preparation The boiling method described by Millar et al. could be used to extract DNA from the bacterial colonies.40 Briefly, one to several colonies from pure cultures are suspended in 50 µL of sterilized water and heated at 100°C for 15 min in a heating block. After centrifugation in a microcentrifuge (6000 × g for 3 min), the supernatant contains bacterial DNA. Alternatively, a variety of DNA extraction kits may be also used.

23.2.2  Detection Procedures 23.2.2.1  16S rRNA Gene Sequencing Principle.  Relman25 utilized primer pair 8FPL (5′-GTTT GATCCTGGCTCAG-3′) and 1492RPL (5′-GGTTACCT TGTTACGACTT-3′) for PCR amplification of the nearly complete length of the 16S rRNA gene. Sequencing analysis of the resulting fragment reveals the identity of the bacteria under investigation. Procedure 1. Prepare PCR mixture (50 µL) consisting of 10 mM Tris-HCl pH 8.3, 50 mM KCl, 1.5 mM MgCl2, 1 mM dNTPs (0.25 mM each), 0.1 µM (each) primer, 1 U of Taq DNA polymerase, and 5 µL (1–5 ng) of template DNA. 2. Conduct PCR amplification with 1 cycle of 94°C for 5 min; 5 cycles of 94°C for 1 min, 52°C for 1 min, and 72°C for 1.5 min; 25 cycles of 94°C for 1 min, 55°C for 1 min, and 72°C for 1.5 min; and a final 72°C for 5 min. 3. Purify and sequence PCR products in both directions by using the above two primers and an additional primer, 1055r (5′-CACGAGCTGACGACAG CCAT-3′).25,27 4. Compare the determined sequences to known sequences of the 16S rRNA genes in the databases of the National Center for Biotechnology Information using the BLASTN algorithm.

252

Note: Species identification is determined from the bestscoring reference sequence of the BLAST output that has a sequence similarity of ≥99% with the query sequence. All positions showing differences from the best-scoring reference sequence could be visually inspected in the electropherogram, and the sequence is corrected if needed, that is, when obvious sequencing software errors occur, or when undetermined nucleotides in the sequence can be determined by visual inspection of the electropherogram. Afterwards, a second search is done with the BLAST algorithm. Ideally, 1300–1500 bp are sequenced with <1% position ambiguities, with a minimum of 500–525 bp being sequenced.24 The criteria for species identification is at least 99% sequence similarity; ideally, >99.5% sequence similarity, with sequence match to type strain or reference strain of species that has undergone DNA-relatedness studies. 23.2.2.2  S equencing of the Ribosomal 16S-23S Intergenic Spacer (ITS) Region Principle.  Gürtler and Stanisich28 designed the bacteriaspecific universal primer pair 13BF (5′-GTGAATACGTTC CCGGGCCT-3′) and 6R (5′-GGGTTYCCCCRTTCR GAAAT-3′) (where Y is C or T and R is A or G) for amplification of a DNA fragment that encompassed ITS, a portion of the 3′ end of the 16S rRNA gene, and a portion of the 5′ end of the 23S rRNA gene.27,30 The 5′ end of primer 13BF is located at position 1371 of the 16S rRNA gene, and the 5′ end of primer 6R is located at position 108 downstream of the 5′ end of the 23S rRNA gene (Escherichia coli numbering). The identity of the bacteria concerned is determined following nucleotide sequencing analysis of the ITS fragment. Procedure 1. Prepare PCR mixture (50 µL) consisting of 10 mM Tris-HCl (pH 8.3), 50 mM KCl, 1.5 mM MgCl2, 0.8 mM deoxyribonucleoside triphosphates (0.2 mM each), 0.1 µM primer (each), 1 U of Taq DNA polymerase, and 5 µL (1–5 ng) of template DNA. 2. Perform PCR amplification with 1 cycle of 94°C for 5 min; 5 cycles 94°C for 1 min; 55°C for 1 min, and 72°C for 1 min; 30 cycles of 94°C for 1 min, 60°C for 1 min, and 72°C for 1 min; and a final 72°C for 3 min. 3. Purify and sequence PCR products in both directions by using the above two primers. Note: Species of Granulicatella produce multiple ITS amplicons in PCR with different length and sequence.27 Only the shortest ITS fragment of a strain, which is usually the dominant band, is eluted from the agarose gel and sequenced. In this way, the ITS of Granulicatella elegans can be sequenced. However, the amplicons of Granulicatella adiacens eluted from gels cannot be directly sequenced due to the presence of different amplicons with similar size in the eluted DNA. For this reason, the eluted DNA from gel can be cloned with a Topo TA cloning kit (Invitrogen, Carlsbad, California, USA). The ITS regions of positive clones are reamplified and sequenced. The determined sequences are compared to

Molecular Detection of Human Bacterial Pathogens

known sequences in the databases of the National Center for Biotechnology Information using the BLASTN algorithm. Species identity is determined from the best-scoring reference sequence of the BLAST output and whether the best-scoring reference sequence in the databases has a sequence identity of ≥98% with the query sequence.27 23.2.2.3  Partial rpoB Gene Sequencing Principle.  Drancourt et al.32 employed primers Strepto F (5′-AARYTIGGMCCTGAAGAAAT-3′) and Strepto R (5′-T GIARTTTRTCATCAACCATGTG-3′) (where M is A or C, R is A or G, and W is A or T) to amplify a portion (740 bp) of the rpoB gene from Streptococcus, Enterococcus, Gemella, Abiotrophia, and Granulicatella. Procedure 1. Prepare PCR mixture (50 µL) consisting of 2.5 U of Taq polymerase, 1× Taq buffer, 1.8 mM MgCl2 (Gibco-BRL/Life Technologies, Cergy Pontoise, France), 0.2 mM concentrations of dATP, dTTP, dGTP, and dCTP, and 0.2 μM concentrations of each primer, and 5 µL (1–5 ng) of template DNA. 2. Subject the PCR mixture to one cycle of 95°C for 2 min; 35 cycles of 94°C for 30 s, 52°C for 30 s, and 72°C for 60 s; and a final 72°C for 5 min. 3. Purify the amplified product by using a QIAquick spin PCR purification kit (Qiagen, Hilden, Germany) and determine the nucleotide sequence of the amplicon. Note: This primer pair Strepto F and Strepto R is shown to be almost specific for streptococcal species, since no amplification products were obtained from 58 other bacterial isolates belonging to nonstreptococcal species, including other grampositive cocci.32 23.2.2.4  Array-Based Method Principle.  Tung et al.39 reported an array-based method for identification of Abiotrophia (1 species), Enterococcus (18 species), Granulicatella (3 species), and Streptococcus (31 species and 6 subspecies). The method consists of PCR amplification of the ribosomal DNA intergenic spacer (ITS) regions, and hybridization of the digoxigenin-labeled PCR products to a panel of 88 oligonucleotide probes (16- to 30-mers) immobilized on a nylon membrane. The probes are designed either from the ITS regions or from the 3′ ends of the 16S rRNA genes, and belong to three categories: species specific, group specific, and supplemental probes. Procedure 1. Amplify the ITS region for hybridization with primer pair 8FPL (5′-GTTTGATCCTGGCTCAG-3′) and 1492RPL (5′-GGTTACCTTGTTACGACTT-3′) as shown in Section 23.2.2.2, except that the reverse primer 6R (5′dig-GGGTTYCCCCRTTCRGAAAT3′) is labeled with a digoxigenin molecule at its 5′ end.39 Labeling of both primers with digoxigenin is encouraged because this can increase the hybri­dization signal.

253

Granulicatella

















2. Fabricate oligonucleotide array: (a) Customer synthesize Granulicatella adiacens– specific probe (5′-TATCACAACAAATAACC AATTAA-3′), Granulicatella elegans probes (5′- GAG GT TA AC TC TCA AC TCGACC T TTGAAAA-3′, 5′-TTTAACAAGAAGCAACG CGACCATA-3′) and Granulicatella balaenopterae-specific probes (5′-ATAAC GGA ACCTACCAAGTTCACTTC-3′, 5′-TGAGAGA T TA AT T C T C T C TAGAC T T T GAT C -3 ′, 5′- G CA A ACG CGA AT CATAT TGAGAC TTAA-3′)30; (b) Add multiple thymine bases (5–15) to the 3′ ends of the above probes to increase the binding of these probes to the nylon membrane that is a solid support for immobilizing probes41; (c) Design a positive control probe (5′-GTCGT AACAAGGTAGCCGTA-3′) from a conserved region at the 3′ end of the 16S rRNA gene (GenBank AB023575) to verify all the procedures including PCR, hybridization, antibody reaction, and colorimetric reaction; (d) Dilute the oligonucleotide probes 1:1 (final concentration, 10 µM) with a tracking dye solution (30% [vol/vol] glycerol, 40% [vol/vol] dimethyl sulfoxide, 1 mM EDTA [disodium salt], 0.15% [wt/vol] bromophenol blue, 10 mM Tris-HCl [pH 7.5]). (e) Draw the probe solutions into different wells of a round-bottom microtiter plate and spotted onto a positively charged nylon membrane (Roche, Mannheim, Germany) with an automatic arrayer (SR-A300; Ezspot, Taipei, Taiwan) by using a solid pin (400–1000 µm in diameter); alternatively, spot the probe solutions on the membrane using the solid pin by hand; use dye solution without probe as a negative control; (f) Dry and expose the membrane to short-wave UV light (Stratalinker 1800, California, USA) for 30 s; Remove unbound oligonucleotides by two washes (2 min each) at room temperature in 0.5 × SSC (1 × SSC is 0.15 M NaCl plus 0.015 M sodium citrate)-0.1% sodium dodecyl sulfate (SDS); (g) Air dry and store the arrays at room temperature under desiccation for further use. 3. Hybridize the labeled PCR products to probes: (a) Prehybridize each array for 2 h with 1 mL of hybridization solution (5 × SSC, 1% [wt/vol] blocking reagent (Roche), 0.1% N-laurylsarcosine,  0.02% SDS) in an individual well of a 24-well cell culture plate. (b) Heat the digoxigenin-labeled PCR product amplified from an isolate in a boiling water bath for 5 min and immediately cool on an ice bath. (c) Hybridize at 45°C for 90 min in an incubator with a shaking speed of 60 rpm.











(d) Wash the array three times (5 min each) in 1 mL of washing buffer (2× SSC, 0.1% SDS) and one wash (1 min) in 1 mL of a second washing buffer (0.5× SSC, 0.1% SDS). (e) Block for 1 h with 1 mL of blocking solution supplied in the DIG Nucleic Acid Detection kit (Roche) (dissolve 1% [wt/vol] blocking reagent in maleic acid buffer [0.1 M maleic acid, 0.15 M NaCl, pH 7.5]). (f) Remove the blocking solution, add 0.5 mL of alkaline phosphatase-conjugated sheep antidigoxigenin antibodies (diluted 1:2500 in blocking solution) to each well, and incubate for 1 h. (g) Wash the array three times (15 min each) in 1 mL of washing solution (0.3% [vol/vol] Tween 20 in maleic acid buffer), wash once in 1 mL of detection buffer (0.1 M Tris-HCl, 0.15 M NaCl, pH 9.5) for 5 min. (h) Add 0.5 mL of alkaline phosphatase substrate (a  stock solution of nitroblue tetrazolium chloride and 5-bromo-4-chloro-3-indolylphosphate diluted 1:50 in detection buffer) (Roche) to each well, and incubate the plate at 37°C in dark (without shaking).

Note: Color development is clearly visible between 30 min and 1 h after the start of the reaction; then the hybridized spot can be easily recognized by the naked eye. A strain is identified as one of the Granlicatella species when the probe (or all probes) specified for that species is hybridized.

23.3  C  ONCLUSION AND FUTURE PERSPECTIVES Members of the genus Granulicatella bacteria are grampositive, catalase-negative, and facultatively anaerobic cocci. Currently, the genus contains three species, Granulicatella adiacens, Granulicatella elegans, and Granulicatella balaenopterae, with the first two being human pathogens. Granulicatella spp. normally display low virulence to hosts and are infrequently isolated as opportunistic pathogens.1 These bacteria may enter the bloodstream and disseminate to other tissues and thus cause infections.8,9 A wide spectrum of infections caused by Granulicatella has been reported.7,10,11,13–17,42,43 They resemble other better known streptococcal isolates and consequently may be misidentified as these bacteria. Granulicatella spp. are part of the normal flora of human pharynx, urogenital, and intestinal tracts.1,7 They form tiny colonies on blood agar supplemented with pyridoxal phosphate or l-cysteine and exhibit satellitism around the colonies of Staphylococcus epidermidis and other streptococci.7 Granulicatella may be overlooked because of their poor growth on unsupplemented media and may not be accurately identified by conventional diagnostic procedures or commercial kits. Molecular techniques are good alternatives for identification of doubtful organisms.

254

Molecular methods used to identify Granulicatella species include 16S rRNA gene sequencing, ribosomal 16S-23S intergenic spacer (ITS) sequencing, partial sequencing of the rpoB gene, and oligonucleotide probe hybridization. Sequencing of the 16S rRNA gene seems to be a prior choice to identify Granulicatella spp. since their respective 16S rRNA gene sequences are available in GenBank. Sequencing of the ribosomal 16S-23S intergenic spacer (ITS) is another alternative. However, Granulicatella spp. possess multiple ITS fragments with different length and sequence; therefore, the amplified ITS should be eluted from agarose gel or cloned before sequencing.30 For this reason, ITS sequencing seems to be impractical for identification of Granulicatella. This drawback can be overcome by the hybridization technique, with probes being designed from the ITS region. Five oligonucleotide probes were successfully used to identify the three Granulicatella species.39 The array format originally developed by Tung et al.39 can be easily transformed to the dot hybridization format without sacrifice of the probe efficacy. Finally, partial sequencing (740 bp) of rpoB, encoding the subunit of the bacterial RNA polymerase, could be used to identify Granulicatella adiacens.32 Unfortunately, there is no rpoB sequence of Granulicatella elegans deposited in public databases. Accurate identification of members of Granulicatella may be of great help in elucidating the epidemiology of these bacteria.

REFERENCES









1. Ruoff, K.L., Aerococcus, Abiotrophia, and other aerobic catalase-negative, gram-positive cocci. In Murray, P. R. et  al. (Editors), Manual of Clinical Microbiology, 9th ed., pp. 443–454, American Society for Microbiology, Washington, DC, 2007. 2. Collins, M.D., and Lawson, P.A., The genus Abiotrophia (Kawamura et al.) is not monophyletic: Proposal of Granulica­ tella gen. nov., Granulicatella adiacens comb. nov., Granulica­ tella elegans comb. nov. and Granulicatella balaenopterae comb. nov., Int. J. Syst. Evol. Microbiol., 50, 365, 2000. 3. Kawamura, Y. et al., Transfer of Streptococcus adjacens and Streptococcus defectivus to Abiotrophia gen. nov. as Abiotrophia adiacens comb. nov. and Abiotrophia defectiva comb. nov., respectively, Int. J. Syst. Bacteriol., 45, 798, 1995. 4. Bouvet, A., Grimont, F., and Grimont, P.A.D., Streptococcus defectivus sp. nov. and Streptococcus adjacens sp. nov., nutritionally variant streptococci from human clinical specimens, Int. J. Syst. Bacteriol., 39, 290, 1989. 5. Roggenkamp, A. et al., Abiotrophia elegans sp. nov., a possible pathogen in patients with culture-negative endocarditis, J. Clin. Microbiol., 36, 100, 1998. 6. Christensen, J.J., and Facklam, R.R., Granulicatella and Abiotrophia species from human clinical specimens, J. Clin. Microbiol., 39, 3520, 2001. 7. Ruoff, K.L., Nutritionally variant streptococci, Clin. Microbiol. Rev., 4, 184, 1991. 8. Coykendall, A.L., Classification and identification of the viridans streptococci, Clin. Microbiol. Rev., 2, 315, 1989. 9. Strom, B.L. et al., Dental and cardiac risk factors for infective endocarditis. A population-based, case-control study, Ann. Intern. Med., 129, 761, 1998.

Molecular Detection of Human Bacterial Pathogens 10. Altay, M. et al., A novel agent of peritoneal dialysis-related peritonitis: Granulicatella adiacens, Perit. Dial. Int., 28, 96, 2008. 11. Biermann, C. et al., Isolation of Abiotrophia adiacens from a brain abscess which developed in a patient after neurosurgery, J. Clin. Microbiol., 37, 769, 1999. 12. Bouvet, A., Human endocarditis due to nutritionally variant streptococci: Streptococcus adjacens and Streptococcus defectivus, Eur. Heart. J., 16 Suppl B, 24, 1995. 13. Cerceo, E. et al., Central nervous system infections due to Abiotrophia and Granulicatella species: An emerging challenge? Diagn. Microbiol. Infect. Dis., 48, 161, 2004. 14. Heath, C.H. et al., Vertebral osteomyelitis and discitis associated with Abiotrophia adiacens (nutritionally variant streptococcus) infection, Aust. N. Z. J. Med., 28, 663, 1998. 15. Hepburn, M.J. et al., Septic arthritis caused by Granulicatella adiacens: Diagnosis by inoculation of synovial fluid into blood culture bottles, Rheumatol. Int., 23, 255, 2003. 16. Namdari, H. et al., Abiotrophia species as a cause of endophthalmitis following cataract extraction, J. Clin. Microbiol., 37, 1564, 1999. 17. Ormerod, L.D. et al., Infectious crystalline keratopathy. Role of nutritionally variant streptococci and other bacterial factors, Ophthalmology, 98, 159, 1991. 18. Sato, S. et al., Abiotrophia elegans strains comprise 8% of the nutritionally variant streptococci isolated from the human mouth, J. Clin. Microbiol., 37, 2553, 1999. 19. Tuohy, M.J. et al., Antimicrobial susceptibility of Abiotrophia adiacens and Abiotrophia defectiva, Diagn. Microbiol. Infect. Dis., 38, 189, 2000. 20. Zheng, X. et al., Antimicrobial susceptibilities of invasive pediatric Abiotrophia and Granulicatella isolates, J. Clin. Microbiol., 42, 4323, 2004. 21. Senn, L. et al., Bloodstream and endovascular infections due to Abiotrophia defectiva and Granulicatella species, BMC Infect. Dis., 6, 9, 2006. 22. Beighton, D. et al., Analysis of enzymatic activities for differentiation of two species of nutritionally variant streptococci, Streptococcus defectivus and Streptococcus adjacens, J. Clin. Microbiol., 33, 1584, 1995. 23. Woo, P.C. et al., Granulicatella adiacens and Abiotrophia defectiva bacteraemia characterized by 16S rRNA gene sequencing, J. Med. Microbiol., 52, 137, 2003. 24. Janda, J.M., and Abbott, S.L., 16S rRNA gene sequencing for bacterial identification in the diagnostic laboratory: Pluses, perils, and pitfalls, J. Clin. Microbiol., 45, 2761, 2007. 25. Relman, D.A., Universal bacterial 16S rDNA amplification and sequencing. In Persing, D.H. et al. (Editors), Diagnostic Molecular Microbiology, pp. 489–495, American Society for Microbiology, Washington, DC, 1993. 26. Chang, H.C. et al., Species-level identification of isolates of the Acinetobacter calcoaceticus–Acinetobacter baumannii complex by sequence analysis of the 16S–23S rRNA gene spacer region, J. Clin. Microbiol., 43, 1632, 2005. 27. Chen, C.C. et al., Identification of clinically relevant viridans group streptococci by sequence analysis of the 16S-23S ribosomal DNA spacer region, J. Clin. Microbiol., 42, 2651, 2004. 28. Gürtler, V., and Stanisich, V.A., New approaches to typing and identification of bacteria using the 16S–23S rDNA spacer region, Microbiology, 142, 3, 1996. 29. Hassan, A.A. et al., Inter- and intraspecies variations of the 16S–23S rDNA intergenic spacer region of various streptococcal species, Syst. Appl. Microbiol., 26, 97, 2003.

Granulicatella 30. Tung, S.K. et al., Identification of species of Abiotrophia, Enterococcus, Granulicatella and Streptococcus by sequence analysis of the ribosomal 16S-23S intergenic spacer region, J. Med. Microbiol., 56, 504, 2007. 31. Mollet, C., Drancourt, M., and Raoult, D., rpoB sequence analysis as a novel basis for bacterial identification, Mol. Microbiol., 26, 1005, 1997. 32. Drancourt, M. et al., rpoB gene sequence-based identification of aerobic gram-positive cocci of the genera Streptococcus, Enterococcus, Gemella, Abiotrophia, and Granulicatella, J. Clin. Microbiol., 42, 497, 2004. 33. Chen, C.C. et al., Identification of clinically relevant viridans streptococci by an oligonucleotide array, J. Clin. Microbiol., 43, 1515, 2005. 34. Su, S.C. et al., Identification of nonfermenting Gram-negative bacteria of clinical importance by an oligonucleotide array, J. Med. Microbiol., 58, 596, 2009. 35. Su, H.-P. et al., Identification of Legionella species by an oligonucleotide array, J. Clin. Microbiol., 47, 1386, 2009. 36. Fukushima, M. et al., Detection and identification of Mycobacterium species isolates by DNA microarray, J. Clin. Microbiol., 41, 2605, 2003. 37. Park, H. et al., Detection and genotyping of Mycobacterium species from clinical isolates and specimens by oligonucleotide array, J. Clin. Microbiol., 43, 1782, 2005.

255 38. Lin, M.C. et al., Identification of six foodborne pathogens and Psedomonas aeruginosa grown on selective media by an oligonucleotide array, J. Food Prot., 68, 2278, 2005. 39. Tung, S.K. et al., Array-based identification of species of the genera Abiotrophia, Enterococcus, Granulicatella, and Streptococcus, J. Clin. Microbiol., 44, 4414, 2006. 40. Millar, B.C. et al., A simple and sensitive method to extract bacterial, yeast and fungal DNA from blood culture material, J. Microbiol. Methods, 42, 139, 2000. 41. Brown, T.J., and Anthony, R.M., The addition of low numbers of 3′ thymine bases can be used to improve the hybridization signal of oligonucleotides for use within arrays on nylon supports, J. Microbiol. Methods, 42, 203, 2000. 42. del Pozo, J.L. et al., Granulicatella adiacens breast implant-associated infection, Diagn. Microbiol. Infect. Dis., 61, 58, 2008. 43. Gephart, J.F., and Washington, J.A. 2nd, Antimicrobial susceptibilities of nutritionally variant streptococci, J. Infect. Dis., 146, 536, 1982. 44. Kanamoto, T. et al., Genetic heterogeneities and phenotypic characteristics of strains of the genus Abiotrophia and proposal of Abiotrophia para-adiacens sp. nov., J. Clin. Microbiol., 38, 492, 2000. 45. Ohara-Nemoto, Y. et al., Infective endocarditis caused by Granulicatella elegans originating in the oral cavity, J. Clin. Microbiol., 43, 1405, 2005.

24 Lactobacillus Ester Sánchez and Yolanda Sanz CONTENTS 24.1 Introduction...................................................................................................................................................................... 257 24.1.1 Taxonomy............................................................................................................................................................. 257 24.1.2 Ecology................................................................................................................................................................. 258 24.1.3 Phenotypic Characterization................................................................................................................................ 258 24.1.4 Genotypic Characterization.................................................................................................................................. 259 24.1.4.1 Molecular Fingerprinting....................................................................................................................... 259 24.1.4.2 PCR and Hybridization Techniques Based on Specific Primers and Probes........................................ 261 24.1.4.3 Sequencing of rRNA and Other Molecular Marker Genes................................................................... 263 24.1.4.4 Temperature and Denaturing Gradient Gel Electrophoresis (TGGE and DGGE)................................ 263 24.2 Methods of the Most Commonly Used Techniques for Lactobacillus Detection............................................................ 264 24.2.1 DNA Sample Preparation..................................................................................................................................... 264 24.2.2 Detection Procedures............................................................................................................................................ 264 24.2.2.1 Identification of Lactobacillus Species by PCR and DNA Sequencing................................................ 264 24.2.2.2 Differentiation of Lactobacillus Species by Multiplex or Simplex PCR............................................... 264 24.2.2.3 Detection and Identification of Lactobacillus Species by DGGE......................................................... 264 24.2.2.4 Quantification of Lactobacillus by Real-Time PCR (SYBR Green)..................................................... 265 24.3 Conclusions....................................................................................................................................................................... 265 Acknowledgments...................................................................................................................................................................... 265 References.................................................................................................................................................................................. 265

24.1  INTRODUCTION The genus Lactobacillus belongs to the lactic acid bacteria (LAB) group, which comprises a broadly defined grampositive bacterial species of nonspore-forming rods or cocobacilli, and which is characterized by the formation of lactic acid as the sole or main end product of carbohydrate metabolism. They are catalase-negative, strictly fermentative, aerotolerant or anaerobic, aciduric or acidophilic, and have complex nutritional requirements and a low GC (guanine + cytosine) content (<50 mol%).1 This genus is one of the most important taxa in food microbiology and human health. Lactobacilli are found in a variety of habitats, where rich carbohydrate-containing substances are available. These include the mucosal membranes of humans and animals (oral cavity, intestine, and vagina), plants and material of plant origin, and man-made habitats such as sewage and fermented or spoiled food. Lactobacillus species play a crucial role in the production and preservation of fermented foods, including vegetables, meats, and dairy products, where they are used as starters, adjunct starters, or protective cultures.2 Lactobacillus species are also found in the human the gastrointestinal tract soon after birth. Some strains are considered probiotics, which confer health benefits

to the consumer, and are commercialized in the form of functional foods or food supplements. Because of the variety of habitats colonized by Lactobacillus species and their impact on food safety, technology, and health, the availability of reliable tools for their detection and identification is of utmost importance. Traditionally, the identification of Lactobacillus strains was based on physiological and metabolic phenotypic traits, such as their carbohydrate fermentation ability. These culture-dependent methods are time consuming and often lead to erroneous results because of the great complexity and phenotypic variability of this genus. Modern developments in molecular biology and increased knowledge of the genome sequences of many bacterial groups have turned molecular methods into the most commonly applied, as they give more rapid and reliable identification results. This chapter includes an overview of the ecology and taxonomy of the Lactobacillus genus and the molecular techniques developed and applied for their detection at species and strain level in complex ecosystems of relevance to human activities.

24.1.1  Taxonomy The genus Lactobacillus was first described by Beijerinck in 1901.3 It belongs to the phylum Firmicutes, class Bacilli, order 257

258

Lactobacillales, and family Lactobacillaceae. The closest relatives are grouped within the same family and include the genera Paralactobacillus and Pediococcus. The phylogenetically closest family appears to be Leuconostocaceae, which includes the genera Leuconostoc, Oenococcus, and Weissella.4 Members of the genus Lactobacillus are gram-positive, nonspore-forming bacteria, and rods or cocobacilli in shape. They are catalase-negative, fermentative, aerotolerant or anaerobic, chemoorganotrophic, aciduric or acidophilic, and have complex nutritional requirements (carbohydrates, amino acids, peptides, fatty acid esters, salts, nucleic acid derivatives and vitamins). Lactobacillus species show a GC content ranging from 32 to 54 mol%. The genus Lactobacillus was initially subdivided into different groups, based on phenotypic characteristics. The first classification was based on their optimal growth temperatures and hexose fermentation pathways.5 Based on 16S rRNA studies, the genus Lactobacillus and related genera were subdivided into three groups: Lactobacillus delbrueckii group, Lactobacillus casei-Pediococcus group, and Leuconostoc group, which also contained some lactobacilli.6 The historical subdivisions of the genus Lactobacillus based on the type of fermentation were reviewed by Hammes and Vogel (1995).1 They divided Lactobacillus species into: (i) obligate homofermentative lactobacilli that ferment hexose sugars almost exclusively (>85%) to lactic acid by the Embden–Meyerhof–Parnas (EMP) pathway, but not pentoses and gluconate because they lack phosphoketolase activity; (ii) facultative heterofermentative lactobacilli that degrade hexoses to lactic acid by the EMP pathway and also ferment pentoses (and often gluconate) because they possess both aldolase and phosphoketolase activities; and (iii) obligately heterofermentative lactobacilli that degrade hexoses by the phosphogluconate pathway producing lactate, ethanol or acetic acid, and carbon dioxide in equimolar amounts; moreover, they ferment pentoses by the same pathway. However, the division of lactobacilli according to the type of fermentation is not in accordance with their phylogenetic clustering as revealed by analysis of their 16S rRNA. The application of polyphasic approaches, which consist of integrating different types of data (phenotypic, genotypic, and phylogenetic) of microorganisms, has been shown to be very useful for taxonomy.7 The use of polyphasic taxonomy has led to taxonomic changes within the Lactobacillus genus. For example, it has led to the reclassification of species, such as L. sobrius strain DSM 16698T as L. amylovorus;8 and genera, such as Pediococcus dextrinicus as L. dextrinicus.9 It has also led to the identification of many new species (e.g., species isolated from sunki, a traditional Japanese pickle: L. kisonensis, L. otakiensis, L. rapi, and L. sunkii).10 At the time of writing (August 2009), in the Approved List of Bacterial Names11 the genus Lactobacillus includes 133 validly described species and 18 subspecies, which reflects its complexity. The description of a large number of new species and the following phylogenetic reexamination of the genus has caused some confusion recently, and earlier lists of identified lactobacilli will continue being modified.12

Molecular Detection of Human Bacterial Pathogens

24.1.2  Ecology Lactobacilli are found in environments where carbohydrates are available, such as food (dairy products, fermented meat, sourdough, vegetables, fruits, and beverages); sewage and plant material; and the respiratory, gastrointestinal, and genital tracts of humans and animals. Lactobacilli are important in the production of foods that involve lactic acid fermentation, notably dairy products (e.g., yogurt and cheese), fermented vegetables (e.g., olives, pickles and sauerkraut), fermented meats (sausages), and sourdough bread. The use of lactobacilli as starter cultures in the food industry has a long history.13 Starter cultures play critical roles in the fermentation processes, development of flavor and texture, and preservation of different fermented foods. For instance, the expression of 15 genes and/or operons was induced in L. sakei during raw-sausage fermentation, reflecting the functional adaptation and roles of this microorganism to the fermentation sausage environment.14 Some Lactobacillus species can also contribute to food spoilage, which are cause of concern in the food industry. The overgrowth of nonstarter species can cause acidification, smear, and out flavors in different products, such as cooked ham and cheeses.15,16 Lactobacillus species are normal residents of the gastrointestinal tract of humans and animals, and have received considerable attention for their possible use as probiotics.17,18 Several recent studies have highlighted the benefits and limitations of the use of specific Lactobacillus strains in different health-related areas, including maintenance of Crohn’s disease remission,19,20 stimulation of the immune system,21,22 prevention of allergic diseases,23 and reduction of the recurrence of Clostridium difficile infections.18,24 For the time being, dairy products are the main vehicle used to supply humans with probiotic strains and, especially, fermented milks, which allow their growth and survival better than other food matrixes. In general, Lactobacillus are avirulent or show low virulence potential; however, members of this genus have been associated with infections in immunosuppressed subjects and patients with health complications, such as bacteremia, endocarditis, and vascular graft infection.25–27 Lactobacilli have also been isolated from carious dentin lesions and are associated with the development of dental caries due to their aciduric characteristics.28

24.1.3  Phenotypic Characterization The phenotypic characterization of lactobacilli usually includes evaluation of morphology, mode of glucose fermentation, growth at different temperatures, configuration of the produced lactic acid, type of fermented carbohydrates, production of methyl esters from fatty acids, and the pattern of whole-cell or cell-wall proteins. However, the use of these traditional methods for the identification of lactobacilli is timeconsuming, requires cultivation, and may lead to erroneous results. In general, phenotypic tests show poor reproducibility

Lactobacillus

and ambiguous results. The expression of phenotypic features depends on the environmental conditions contributing to the variability of the results; for example, growth at different temperatures changes the lipid composition of the membrane of L. acidophilus CRL 640.29 Moreover, lactobacilli share many biochemical and phenotypical characteristics within species of the same genus and other related genera. Altogether, these drawbacks adversely affect the reliability of phenotype-based methods as unique tools for identification of lactobacilli at genus and at species level. Commercial kits based on biochemical and enzymatic analysis has been used for long time as simple and fast methods for the identification of Lactobacillus at species level. Nevertheless, the results of these analyses, for instance using the API 50 CH strips, did not have good reproducibility, probably due to their susceptibility to different environmental and physiological conditions. These data suggest that the use of the current API 50 CH database and other similar methods for identification of Lactobacillus species could lead to misidentification, when applied as the only identification tool.30 Anyway, phenotypic characterization is important for a preliminary classification as well as to get basic knowledge of the properties of Lactobacillus strains. Salt and pH tolerance, growth at certain temperatures, configuration of the produced lactic acid, antibiotic resistance, and bacteriocin production are important characteristics for selecting Lactobacillus strains as possible starter or protective cultures and as probiotics.

24.1.4  Genotypic Characterization Many different genotyping techniques can be applied for species identification, strain differentiation, and quantification. The major advantages of these DNA-based typing methods lie in their discriminatory power, universal applicability, and culture independency.31 Species and closely related strains with similar phenotypic features can be distinguished by DNA-based molecular fingerprinting techniques, including restriction fragment length polymorphism (RFLP), pulse field gel electrophoresis (PFGE), randomly amplified polymorphic DNA (RAPD), and ribotyping: identification and quantification of species and groups can be achieved by molecular techniques based on the use of specific primers and probes, including conventional polymerase chain reaction (PCR), real-time PCR hybridization techniques, and sequencing of rRNA and other gene markers. Moreover, detection and evolution of species in complex ecosystems can be achieved by denaturing gradient gel electrophoresis (DGGE) and temperature gradient gel electrophoresis (TGGE). These methodologies and examples of their application to the genus Lactobacillus are described below. 24.1.4.1  Molecular Fingerprinting Many common typing methods have been adapted to the genus Lactobacillus. These techniques are applied to differentiate between species and strains, which may belong to the same species, previously isolated on a culture medium. The

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techniques used for the fingerprinting of Lactobacillus species are summarized in Figure 24.1. Restriction Fragment Length Polymorphism. In RFLP analysis, the DNA is digested by restriction enzymes and the resulting restriction fragments are separated according to their lengths by gel electrophoresis. The banding patterns that result are referred as DNA fingerprinting. Because of the high specificity of restriction enzymes and the stability of chromosomal DNA, a reproducible pattern of fragments is obtained after the complete digestion of the chromosomal DNA by a particular enzyme. One general criticism about this method is the complexity of banding patterns. However, this technique can be applied on a selected genetic marker by its amplification with specific primers (PCR–RFLP), reducing its complexity. Then, the generated DNA fingerprinting and the discrimination power between different bacterial species or subspecies, will be dependent on the restriction enzymes used and the length of the amplified fragment. Amplified ribosomal DNA restriction analysis (ARDRA) is essentially PCR–RFLP analysis of the amplified 16S rRNA gene. ARDRA patterns are highly reproducible and comparable between laboratories, but the discriminatory power of this technique is generally low because of the conserved nature of this gene. Other highly conserved genes, such as rpoB, hsp60, or the internal spacer region between the 16S and 23S rRNA genes (ISR), and their restriction by different enzymes have been used for Lactobacillus identification by PCR–RFPL. Chenoll et al. (2006)32 identified L. tucceti as a novel specie based on their singular ISR-DdeI and ISRHaeIII profiles that allowed its differentiation from 68 lactic acid bacteria reference strains and showed that the discriminatory power of the ISR digestion was better than that of the 16S rRNA gene.33 Furthermore, RFLP analysis of the rpoB sequence amplified by PCR has successfully been used to differentiate 12 lactobacilli at special level after two or three digestions (AciI, HinfI, and MseI).32 Pulse Field Gel Electrophoresis. PFGE employs an alternating field of electrophoresis to allow the separation of large DNA fragments obtained from restriction digests of genomic DNA with rare-cutting enzymes (i.e., SfiI, NotI, ApaI, SgrA, and SmaI). PFGE is performed as a standard gel electrophoresis, but the voltage is periodically switched among three directions; one that runs through the central axis of the gel, and two that run at an angle of 120 degrees either side.34 The technique can be more time consuming than other fingerprinting strategies (4–5 days). However, the profile generated by PFGE has a very good discriminatory power. Indeed, PFGE technique has been successfully used for Lactobacillus species and subspecies differentiation.34–36 For example, the diversity and dynamics of Lactobacillus populations were monitored throughout the manufacturing of Camembert cheese. L. paracasei was the species most frequently isolated and these isolates (39) were classified by PFGE into 21 different profiles among the same species.37 The typing efficacy of this technique can be increased by the use of several single restriction enzymes, followed by numerical analysis of the combined patterns. Sánchez et al (2004).35

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Molecular Detection of Human Bacterial Pathogens

(a) RFLP

1

2

3

4

2 3 1

DNA

Restricted DNA with endonucleases

4 Agarose gel A+

(b) PFGE

B–

B+ Cells embedded in agarose

Enzimatic purification of DNA

A– Size separate DNA fragments

(c) RAPD 1

3 4

2

3

1

4 Targeted DNA

2 Agarose gel

(d) Ribotyping

DNA

Restricted DNA with endonucleases

Size separate DNA fragments

Hybridized whith rDNA probes

FIGURE 24.1  Fingerprinting techniques for Lactobacillus species and strains (a) Restriction fragment length polymorphism (RFLP). The DNA fragment is restricted into pieces (for example 1–4) by a restriction enzyme. The restricted DNA is then loaded into a well in an agarose gel and the fragments are separated by electrophoresis, based on fragment size. (b) Pulse field gel electrophoresis (PFGE). Whole cells are embedded in agarose and DNA is enzymatically purified and restricted by rare-cutting enzymes in situ. The agarose plug is then inserted into a well in an agarose gel and the large restricted fragments are separated by an electric current which pulses from different angles. (c) Random amplification of polymorphic DNA (RAPD). PCR is run with arbitrary sequence primers that amplify segments of DNA of unknown identity. The amplified DNA is then loaded into an agarose gel and the fragments are separated by electrophoresis, based on fragment size. (d) Ribotyping. DNA is isolated from a cultured isolate, restricted and size separated in an agarose gel. The gel is then hybridized with a labeled rRNA/rDNA probe, which binds to fragments containing copies of the specific rRNA. Following probe detection, fragments with bound probe are visualized.

used five rare-cutting enzymes (SfiI, NotI, ApaI, SgrA, and SmaI) for PFGE analysis of 149 lactobacilli isolated from fermented vegetables. The use of SfiI and SmaI led to the most informative digestion patterns, and their combined numerical analysis improved the discriminatory power of the technique.35 Some comparative studies of different typing techniques have indicated that isolates that are indistinguishable by PFGE are unlikely to demonstrate substantial differences by other typing techniques.38,39 Nevertheless, PFGE patterns can be used for bacterial identification only when a set of known strains is included in the same analysis and when a suitable and objective method of analysis of the patterns is applied.40

Random Amplification of Polymorphic DNA Finger­ printing. The RAPD technique is a PCR-based discrimination method in which short arbitrary primers (8–12 nucleotides) anneal to multiple random target sequences of the genomic DNA, resulting in patterns of diagnostic value. Low-stringency annealing conditions are used in PCR reactions, which results in the amplification of randomly sized DNA fragments. RAPD–PCR is a molecular-typing method that is very suitable for identification and differentiation of Lactobacillus strains. It is less time consuming than PFGE, but the construction of a database is nearly impossible due to the high variability of the results. If an unidentified strain has

Lactobacillus

to be investigated a set of known strains has to be included in the same RAPD procedure. As the reproducibility of RAPD patterns is occasionally poor, this method needs to be performed under carefully controlled conditions. RAPD technique has been reported as a very useful tool for Lactobacillus differentiation at strain level,15,16,41,42 but other complementary tools are required for species identification, such as sequencing of molecular marker genes, DGGE, or TGGE. In this context, for example, Dimitonova et al. (2008), compared the RAPD patterns of 20 vaginal lactobacilli isolates, generated with the M13V primer. Two main clusters were generated and divided into subgroups. Identical RAPD patterns were shown in different isolates, suggesting that they were multiple isolates from a single species and identical strains. ARDRA-HaeIII analysis and 16S rRNA gene sequencing allowed the identification at species level of seven strains, three as L. fermentum, two as L. gasseri, one as L. brevis, and one as L. salivarius.43 Ribotyping. A ribotype is essentially a RFLP profile consisting of the restriction fragments from a particular genome, which contain rRNA genes. The bacterial chromosomal DNA is totally restricted into multiple fragments, using a specific restriction enzyme. The restricted fragments are then separated by agarose gel electrophoresis and hybridized with 16S and 23S rRNA probes. The hybridization can be carried out directly in the gel using gel hybridization techniques or on a nylon or nitrocellulose membrane following Southern transfer DNA from the gel to the membrane.44 Following probe detection, restriction bands containing copies of the rRNA genes are visualized and the pattern of the band sizes represents a characteristic fingerprint. Ribotyping fingerprint patterns are more stable and more easily interpretable than those obtained by only restriction enzyme analysis; furthermore, they are highly reproducible, and a single rRNA probe can be used for typing all bacteria because of the similarity of ribosomal genes.45 Ribotyping technique also allows the differentiation of Lactobacillus isolates at strain level. In this context, for example, the probiotic strain L. rhamnosus 35 was characterized at molecular level and compared with seven reference strains from the L. casei group by ribotyping.46 This molecular technique is also a useful tool for the identification of new species, for instance, Lactobacillus similis.47 Automated ribotyping has also been used for the characterization of Lactobacillus. For instance, 91 type and reference stains were classified in different ribotypes. Strains of the L. casei group were divided into five ribotyping genotypes, and strains of the L. acidophilus group were divided into two major clusters, one containing L. acidophilus, L. amylovorus, L. crispatus and L. gallinarum, and another containing L. gasseri and L. johnsonii.48 Several studies have compared the discriminatory power of different molecular-typing methods.35,49,50 In general, the most commonly applied techniques can be placed in the following order with respect to their discriminatory power: PFGE > ribotyping > RAPD > ARDRA. The choice of the most appropriate typing method also depends on other

261

features such as cost, high throughput capacity, and reproducibility of the fingerprints. 24.1.4.2  P  CR and Hybridization Techniques Based on Specific Primers and Probes Specific oligonucleotide probes and primers targeting rRNA or rDNA have been designed for detection and quantification of different species of the Lactobacillus genus using diverse PCR and hybridization techniques. Lactobacilli are phylogenetically heterogeneous, which causes difficulties in the design of a very specific genus probe, but some group-specific probes and primers have been described.51 Frequently, Lactobacillus group-specific primers cover, in addition to Lactobacillus, Leuconostoc, Pediococcus, and Weissella genera.52,53 Speciesspecific probes and primers have also been designed for the identification of Lactobacillus species, but the probe panel is still incomplete, and the detection of some species by these techniques is not possible yet.51 Conventional and Real-Time PCR. Amplification of DNA by PCR is one of the easiest and most rapid ways of detecting and identifying specific bacterial sequences with the use of adequate primers. A summary of primers and conditions designed for the detection of members of the Lactobacillus genus is shown in Table 24.1. The conventional PCR is sufficiently sensitive to detect Lactobacillus at genus57,58 and at specie level.52–56 More recently, multiple PCR approaches have been designed to detect simultaneously different species in a single PCR reaction. For example, it has been possible to simultaneously distinguish L.  plantarum, L. pentosus and L. paraplantarum species by using a multiplex PCR protocol. The size of the amplicons was 318 bp for L. plantarum, 218 bp for L. pentosus, and 107 bp for L. paraplantarum.59 Conventional PCR can be used for semiquantitative analyses by making dilutions of known amounts of DNA almost identical to the target.60 Real-time PCR is a DNA-based technique that monitors the amplification of the target DNA in real time by fluorescence detection. The amount of DNA corresponding to gene copy numbers or bacterial cells in a sample is determined by measuring the cycle number at which the increase in fluorescence (and therefore cDNA) is exponential. Real-time PCR can be used to quantify bacteria from various samples including milk, feces, food, and water. Contemporary, realtime PCR allows the monitoring of the complete amplification and a more accurate quantification. Even though, the determination of bacterial numbers is difficult due to factors affecting the amplification reaction and variation of the copy number and the ribosomal content of cells in different bacteria.61 In spite of these limitations, sets of species-specific 16S rRNA primers and real-time PCR have been used successfully for monitoring and quantifying, for example, the genus Lactobacillus in feces of patients with celiac disease and healthy controls.62,63 Scheirlinck et al. (2009)64 also studied the predominant sourdough LAB species in the production of sourdough bakeries. Real-time PCR assays were developed using species-specific primers targeting the pheS gene of L. plantarum and L. sanfranciscensis to detect, quantify, and monitor them in sourdough and bakery environments.64

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Molecular Detection of Human Bacterial Pathogens

TABLE 24.1 16S rRNA Targeted PCR Primers Designed for the Detection of Lactobacillus Species Specificity L. acidophilus

L. casei

L. casei/paracasei L. casei/paracasei/ rhamnosus/zeae L. crispatus

L. delbrueckii

L. fermentum L. gallinarum L. gasseri L. johnsonii

L. paracasei

L. plantarum L. reuteri L. rhamnosus

L. salivarius

Sequence (5′–3′) GATCGCATGATCAGCTTATA AGTCTCTCAACTCGGCTATG AGCTGAACCAACAGATTCAC ACTACCAGGGTATCTAATCC GCGGACGGGTGAGTAACACG GCTTACGCCATCTTTCAGCCAA GCGGAAATAGTAGTGTGACGATC GCGGAAATAGGTGCATAGGCG CTCAAAGCCGTGACGGTC ACGTGGTGCTAATAATCCTAGTG GTAATGACGTTAGGAAAGCG ACTACCAGGGTATCTAATCC GTGCTTGCACTGAGATTCGACTTA TGCGGTTCTTGGATCTATGCG TCTTGATTTAATTTTG GCACCGAGATTCAACATGG GCGGACGGGTGAGTAACACG GCTTACGCCATCTTTCAGCCAA GGTTCTTGGATYTATGCGGTATTAG ACATGAATCGCATGATTCAAG AACTCGGCTACGCATCATTG AGCGAGCGGAACTAACAGATTTAC GCGGACGGGTGAGTAACACG GCTTACGCCATCTTTCAGCCAA GGRTGATTTGTTGGACGCTAG GCCGCCTTTCAAACTTGAATC GCACCTGATTGATTTTGGTCG GTTGTTCGCATGAACAACGCTTAA CGGTAATGACGCTGGGGAC CAGTTACTACCTCTATCTTTCTTCACTAC GAGCTTGCCTAGATGATTTTA ACTACCAGGGTATCTAATCC CACTAGACGCATGTCTAGAG AGTCTCTCAACTCGGCTAT GTGCTTGCACCGAGATTCAACATG TGCGGTTCTTGGATCTATGCG CCGAGATTCAACATGG ATCATGATTTACATTTGAGTG TGAATTGACGATGGATCACCAGTG TGCTTGCATCTTGATTTAATTTTG GTGCTTGCATCTTGATTTAATTTT TGCGGTTCTTGGATCTATGCG CGAAACTTTCTTACACCGAATGC ATTCACTCGTAAGAAGT

Hybridization Techniques. Hybridization techniques with specific probes are also useful for the detection and quantification of lactobacilli. Of these techniques, the most commonly used are dot blot hybridization and fluorescence in situ hybridization (FISH). Dot blot hybridization is used for the identification of specific nucleic acid sequences or genes with varying degree of homology and for the estimation of relative amounts of nucleic

Target Site

Reference

205–224 328–309 70–89

52 53 53 54 54 53

61–84 185–165 68 86–104 93–112 213–192 196–169

52

65–89 185–205 305–286 92–102 219–199 86–94 160 78–97 470–442

56 52

170–189 296–277 60–83 185–165 69 96 91 81–104 88–111 213–194 67–91 95

52

55 56 52 56 52

56 56 56 55 56 56 53

52 55 55 55 56 52 56 55

acid with known homology. DNA or RNA is extracted from a sample or cultured bacteria, fixed to a nylon or nitrocellulose membrane and hybridized with a labeled oligonucleotide or DNA fragment complementary to the nucleic acids of interest. For example, dot blot hybridization analysis has been used for screening the presence of Lactobacillus reuteri and the gene encoding for the enzyme responsible for reuterin production (gldC).65

Lactobacillus

In FISH, the detection of rRNA sequences within morphologically intact cells is achieved using fluorescently labeled oligonucleotide probes. Special permeabilization procedures may be required to aid passage of the probe into the cells of some bacteria. The multiple rRNA molecules present in bacterial cells capture the probe, the signal is intensified, and the cells can be detected and numerated by using an epifluorescence microscope or flow cytometry.66,67 For example, using fluorescent L. plantarum-specific targeted oligonucleotide probes, the presence of L. plantarum L2 on the jejunum, ileum and colon of rats was demonstrated, after its oral administration.68 The use of FISH coupled with flow cytometry analysis has also demonstrated that Lactobacillus population is significantly reduced in the intestinal microbiota of patients with celiac disease compared with healthy children.69 24.1.4.3  S equencing of rRNA and Other Molecular Marker Genes Relatedness among organisms is estimated through the comparison of molecular sequences, mainly 16S rRNA encoding genes.70 It is based on several major assumptions: rRNA genes are highly conserved because of the fundamental role of ribosomes in the protein biosynthesis that was developed in the early stages of the evolution of microorganisms. Horizontal gene transfer phenomena among organisms have not involved those genes, and the amount of similarity of these sequences between different individuals is representative of the variation of their genomes. The 16S rRNA gene is a well-conserved universal marker, but there are some shortcomings associated with its use. First, the 16S rRNA genes are so well conserved that it has limited resolving power. Second, although the 16S rRNA gene is a universal marker, different bacterial species have different copy numbers of the gene and this leads to an over- and underrepresentation of some bacterial species. In general, if the 16S rRNA gene sequence identity of two microorganisms is lower than 97%, they are poorly related at the genomic level and, therefore, they are considered to belong to different species. If the 16S rRNA gene sequence identity of two microorganisms is higher than 97%, even if they are identical sequences, they have to be considered closely related, and only total DNA–DNA hybridization data and/or analysis of other more discriminative gene sequences are decisive for the identification at the species level. Phylogenetic and alignment analyses are largely preferred over DNA hybridization tests because the latter are time consuming, complex, and expensive. In contrast, identification through sequence analysis is based on specific DNA amplification, sequencing reaction, and comparison with public databases, which are very fast, more reproducible and much less expensive.12 In general, 16S rRNA gene analysis has been used for the identification of different Lactobacillus species,71,72 but the high similarities of the 16S rRNA gene in this bacterial group usually require the confirmation by the analysis of other markers. Thus, the sequencing of several other genes, such as the recA (encoding recombinase A)42,61,73; tuf gene (encoding

263

elongation factor Tu, involved in protein biosynthesis)74,75; hsp60 (groEL, encoding a 60-kDa heat shock protein)76; mal (encoding malolactic enzyme)77; and rpoB (encoding RNA polymerase beta subunit)32 has been used for the identification of lactobacilli. 24.1.4.4  T  emperature and Denaturing Gradient Gel Electrophoresis (TGGE and DGGE) For DGGE and TGGE analysis, RNA or DNA is extracted from an environmental sample and is amplified with specific primers. In DGGE as well as in TGGE, DNA fragments of the same length, but with different sequences, can be separated by electrophoresis through a polyacrylamide gel containing a linear gradient of DNA denaturants formed by urea and formamide (DGGE) or a linear temperature gradient (TGGE). Separation is based on the melting of rRNA genes at specific denaturing points based on their sequence; therefore, each individual sequence will begin to melt at a characteristic denaturing point. The melting changes the conformation of the DNA molecule, slowing its migration though the gel. By using DGGE or TGGE, 50% of the sequence variants can be detected in DNA fragments up to 500 bp. This percentage can be increased to nearly 100% by the attachment of a GC-rich sequence to one side of the DNA fragment. A GC clamp is generally included at the five-end of one of the PCR primers, co-amplified, and thus introduced into the amplified DNA fragments in order to maintain the double-stranded integrity of the amplified products, which improves the detection. The gene sequences from bacterial species in a mixed culture are first amplified using conserved bacterial primers (universal or group-specific primers) that target a hypervariable region, producing amplicons of the same length but with differing sequences that are specific at specie level. DGGE and TGGE allow the separation of these amplicons, producing a molecular fingerprint of the bacterial species present in a sample. The design of primers based on sequence variability in 16S rRNA, 23S rRNA, and other molecular marker genes together with DGGE and TGGE techniques have improved our understanding of the lactobacilli communities present in a variety of complex ecosystems, including the intestinal tract52,78,79 and fermented food.32,42 DGGE and TGGE techniques are useful tools for monitoring both dynamic changes in mixed populations over time and the diversity of Lactobacillus communities because they are rapid and economical. However, they also have several limitations.80 First, only relatively small fragments can be separated by DGGE and TGGE, limiting the amount of sequence information for phylogenetic inferences. Second, the use of different regions of the 16S rRNA and different DGGE and TGGE conditions might result in different resolutions of separation. Finally, DGGE and TGGE only display the DNA fragments obtained from the predominant species present in the community. Several studies revealed that lactobacilli make up less than 1% of the gastrointestinal community, and in this case, the use of a nested approach improves the detection of bands in the denaturing gradient gels. The nested-PCR approach

264

consisting in a first amplification with universal bacterial primers and a second amplification with specific primers for Lactobacillus and closely related species.81

24.2  M  ETHODS OF THE MOST COMMONLY USED TECHNIQUES FOR LACTOBACILLUS DETECTION 24.2.1  DNA Sample Preparation Lactobacillus DNA can be extracted with conventional DNA extraction procedures using lysozyme, proteinase K, and sodium dodecyl sulfate, with slight modifications depending on the type of starting material.82 Commercialized kits may also be used for genomic DNA extraction from fresh or frozen human stool or other sample types, using a rapid and simplified procedure. DNA extracts from reference bacterial strains can also be obtained from a high, concentrated cell suspension in 50 µL of sterile water by boiling for 10 min and freezing for 5 min at −20°C. To verify whether a particular DNA extraction method is effective for a specific matrix, it should be tested on samples containing a well-known number of lactobacilli cells.

24.2.2  Detection Procedures 24.2.2.1  I dentification of Lactobacillus Species by PCR and DNA Sequencing Principle.  Lactobacillus spp. can be identified by PCR amplification and sequencing analysis of 16S rRNA using the universal primers 27f and 1492R. The primer sequences are: 27f, 5′-AGAGTTTGATCCTGGCTCAG-3′ 83; and 1495R, 5′-CTACGGCTACCTTGTTACGA-3′.84 Procedure.  The reaction mixture (50-µL) is performed as follows: 5 µL of extracted DNA, 5 µL 10× buffer, 0.2 mM of dNTP, 2.5 mM MgCl2, 10 pmol of each primer, and 1 U of Taq DNA polymerase. Amplification is done using a PCR System programmed as follows: 94°C for 5 min; 30 cycles of 94°C for 1 min, 58°C for 1 min and 72°C for 2 min; and finally, 72°C for 10 min. PCR products (1.5 kb) are electrophoresed on a 1% agarose gel in 1× Tris-Borate-EDTA, and visualize by UV transillumination after ethidium bromide staining. Sequence the PCR products by using the forward primer and the reverse primer. For bacterial identification, partial 16S rRNA gene sequences are compared with sequences deposited in the GenBank database, using the program standard nucleotidenucleotide Basic Local Alignment Search Tool (BLAST). A threshold of 97% similarity in 16S rRNA gene sequence has been proposed for the bacterial-species delineation based on a correlation study between DNA–DNA hybridization results and similarity values for 16S rRNA gene sequences. 24.2.2.2  D  ifferentiation of Lactobacillus Species by Multiplex or Simplex PCR Principle.  The species Lactobacillus plantarum, Lactobacillus pentosus, and Lactobacillus paraplantarum can be

Molecular Detection of Human Bacterial Pathogens

differentiated by doing a unique PCR reaction with recA derived primers. The multiplex PCR assay can be performed with the primers paraF (5′-GTCACAGGCATTACGAAAAC-3′), pentF (5′-CAGTGG CGCGGTTGATATC-3′), planF (5′-CC GTTTATGCGGAACACCTA-3′), and pREV (5′-TCGGG ATTACCAAACATCAC-3′).59 Lactobacillus species can also be detected by simplex PCR using primers specifics for each specie in each reaction, as those summarized in Table 24.1. Procedure.  The PCR mixture (20 µL) is composed of 5 µL of DNA, 2 µL 10× buffer, 1.5 mM MgCl2, 0.12 mM of the primer planF and 0.25 mM of the primers paraF, pentF, and pREV, 0.3 mM of dNTPs, and 1 U of Taq DNA polymerase. PCRs are performed using a PCR System programmed as follows: 94°C for 3 min, 30 cycles of 94°C for 30 s, 56°C for 10 s, and 72°C for 30 s; and final extension at 72°C for 5 min. The PCR products are then separated electrophoretically on an agarose gel and stained with ethidium bromide. L. plantarum, L. pentosus, and L. paraplantarum strains generate a 318-, 218-, and 107-bp fragments, respectively. Similarly, the amplification products of a simple PCR reaction can be detected on an agarose gel and identified according to its expected size by ethidium bromide staining. 24.2.2.3  D  etection and Identification of Lactobacillus Species by DGGE Principle.  Fecal Lactobacillus communities can be monitored by PCR amplification and DGGE analysis of using 16S rRNA group specific primers. Procedure.  Genomic DNA is amplified using the primers Lac1 (5′-AGCAGTAGGGAATCTTCCA-3′); and Lac2 (5′-ATTYCACCGCTACACATG-3′). A GC clamp (5′-CGCCCGGGGCGCGCCCCGGGCGGCCCGGGG GCACCGGGGG-3′) was attached to the reverse primer (Lac2) to obtain PCR fragments suitable for DGGE analysis.52 Amplification can be carried out using a GeneAmp 2400 Thermocycler (Perkin-Elmer). The reaction mixture (50 μL) contained 25-pmol amounts of each primer, 0.2 mM of each dNTP, reaction buffer, 2 mM MgCl2, 25 μg of bovine serum albumin, 2.5 U of Taq polymerase (Amersham Pharmacia Biotech), and 1 μL of DNA solution. The amplification program is 94°C for 2 min; 35 cycles of 94°C for 30 s, 61°C for 1 min, and 68°C for 1 min; and finally, 68°C for 7 min. DGGE analysis of PCR amplicons is carried out on the Dcode Universal Mutation Detection System (Bio-Rad, Richmond, CA). The denaturing gradient of urea and formamide used for PCR products separation is 25%–45%. A 100% denaturant corresponds to 7M urea and 40% (v/v) formamide. DGGE ladders for the identification of Lactobacillus-like populations are prepared by mixing equal amounts of amplicons obtained from selected reference Lactobacillus species using primers Lac1 and Lac2-GC. Selected unknown DGGE bands can be excised from the acrylamide gels and reamplified with primers Lac1 and Lac2. After purification, sequencing is caring by DNA sequencing using an ABI PRISM-3130XL Genetic Analyzer (Applied

265

Lactobacillus

Biosystems, California, USA). Alternatively isolated bands can be cloned into a vector, checked by PCR, and sequenced afterwards. Sequences identities are determined in GenBank database. 24.2.2.4  Q  uantification of Lactobacillus by Real-Time PCR (SYBR Green) Principle.  Genomic DNA has been used to detect and quantify Lactobacillus spp. with high sensibility by using group-specific primers and real-time PCR (SYBR Green). The primer­ ssequences used are: 5′-AGCAGTAGGGAATCTTCCA-3′, and 5′-CACCGCTACACATGGAG-3′.85 Procedure.  Quantitative PCR can be performed with an iCycler iQ apparatus (Bio-Rad, USA) associated with the iCycler optical system interface software (version 2.3; Bio-Rad) or other equivalent equipments. All PCRs are performed in a volume of 25 µL, using 96-well optical grade PCR plates and an optical sealing tape (Bio-Rad). Reaction mixtures for the optimized SYBR Green assays consist of 1:75,000 dilution of SYBR Green (Molecular Probes); 10 mM Tris-HCl (pH 8.8); 150 mM KCl; 0.1% Triton X-100; 2 mM MgCl2; 100 µM each dNTP; 0.5 µM of each primer and 0.02 U Dynazyme II µL−1 (Finnzymes, Finland); and either 5 µL template or water (no-template control). The thermal cycling conditions are an initial DNA denaturation step at 95°C for 5 min, followed by 35 cycles of denaturation at 95°C for 15 s; primer annealing at 58°C for 20 s; extension at 72°C for 30 s, and an additional incubation step at 80°C–85°C for 30 s to measure the SYBR Green fluorescence. Finally, melt curve analysis was performed by slowly cooling the PCRs from 95°C to 60°C (0.3°C per cycle) with simultaneous measurement of the SYBR Green signal intensity. The bacterial concentration from each sample is calculated by comparing the Ct (cycle threshold) values obtained from standard curves. Standard curves can be created by using serial tenfold dilution of pure culture DNA corresponding to 102–109 cells.

24.3  CONCLUSIONS This decade has seen the emergence of numerous molecular approaches for the analysis of diverse aspects of Lactobacillus populations in complex ecosystems, including the gastrointestinal tract and different foods, and for the identification of new species. Quantitative and qualitative detection can be optimally achieved by the combined use of different methods that complement each other. Thus, numbers can be quantified by using labeled group-specific probes and fluorescent in situ hybridization (FISH), coupled with either microscopy or flow cytometry detection, or by real-time PCR. The species diversity can be subsequently determined by PCR in combination with DGGE, TGGE or DNA sequencing. Strain differentiation could be possible by using PFGE, ribotyping, and RAPD. The application of these techniques is contributing to a comprehensive analysis of the ecological distribution, functional roles, and adaptation of Lactobacillus strains to different

environments. In the near future, it can be anticipated that the increasing use of massive sequencing approaches to the analysis of whole bacterial genomes, metagenomics, and transcripomics will lead to the identification of novel species and roles of Lactobacillus strains in human health and food safety and technology.

ACKNOWLEDGMENTS This work was supported by grants AGL2007–66126-C03–01/ ALI and Consolider Fun-C-Food CSD2007–00063 from the Spanish Ministry of Science and Education (MEC). The scholarship to E. Sánchez from Institute Danone is fully acknowledged.

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25 Leuconostoc María Mar Tomás and Germán Bou CONTENTS 25.1 Introduction...................................................................................................................................................................... 269 25.1.1 Taxonomy of Genus Leuconostoc........................................................................................................................ 269 25.1.2 Isolation and Identification of Leuconostoc spp................................................................................................... 269 25.1.3 Virulence and Clinical Significance..................................................................................................................... 272 25.1.4 Antibiotic Susceptibilities..................................................................................................................................... 272 25.1.5 Molecular Identification of Leuconostoc spp....................................................................................................... 272 25.2 Methods............................................................................................................................................................................ 274 25.2.1 Sample Preparation............................................................................................................................................... 274 25.2.2 Detection Procedures............................................................................................................................................ 275 25.3 Conclusion........................................................................................................................................................................ 275 References.................................................................................................................................................................................. 275

25.1  INTRODUCTION Members of the genus Leuconostoc (leucus, clear, light; nostoc, algal generic name; leuconostoc, colorless nostoc) are facultatively anaerobic, catalase-negative, gram-positive cocci organized in pairs and chains.1 They are heterofermentative lactic acid bacteria found naturally in milk, grass, herbage, grapes, many vegetables, meat, and sugar.2,3 Members of this group are used in dairy fermentations to produce aromatic compounds.4

25.1.1  Taxonomy of Genus Leuconostoc Species of Leuconostoc are the only catalase-negative cocci that are vancomycin resistant, PYR and LAP negative, and that produce CO2 from glucose. They could appear morphologically similar to:

1. Genus Weissella, which includes organisms previously classified as Leuconostocs and the species formerly named Lactobacillus confusus. 2. Alpha-hemolytic or nonhemolytic Enterococcus or Lactococcus spp. It happens when these bacteria grown on the surface of 5% sheep blood agar plates. In the past it was known that Leuconostoc spp. was associated with human diseases5 because these bacteria were generally identified as variants of established human pathogens, such as viridans streptococci or enterococci, or more frequently, were reported as unidentified gram-positive cocci. As many as 31% of Leuconostoc spp. react with the group D antiserum and may be reported as



unidentified Enterococcus spp. The nonserogroup D bacteria have characteristics that sometimes lead to their identification as Streptococcus uberis, S. bovis, S. sanguis, S. equinus, S. anginosus group, or Aerococcus-like or Lactobacillus-like bacteria. 3. The heterofermentative vancomycin-­resistant Lactobacillus spp. However, species of Leuconostoc spp. tend to be more coccoid-like than the Lactobacillus spp. Cautious interpretation of the Gram stain coloring from growth of these bacteria in thioglycolate broth is important for differentiation of leuconostocs and gas-forming lactobacilli.

With the recognition of Leuconostoc spp. as human pathogens, such mistakes in identification are now odd because of the unique results produced by leuconostocs in the tests listed in Table 25.1.6

25.1.2  Isolation and Identification of Leuconostoc spp. Isolation Procedure. Generally, there are no special requirements for isolation of the group of bacteria discussed here; general recommendations for the culture of blood, body fluids, and other specimens should be followed. These organisms are likely to be isolated on rich, nonselective media (e.g., blood and chocolate agars and thioglycolate broth) since they are nutritionally fastidious. If selective isolation of members of the vancomycin-resistant genera Leuconostoc and Pediococcus is desired, Thayer–Martin medium may be used to inhibit normal flora or other contaminating microorganisms.34 269

270

Molecular Detection of Human Bacterial Pathogens

TABLE 25.1 Basic Phenotypic Characteristics of Catalase-Negative, Gram-Positive Cocci Organisms (reference[s])

PYR

LAP

NaCl

BE

Abiotrophia defectiva7–9 Aerococcus christensenii10,11 Aerococcus sanguinola11,12 Aerococcus urinae11,13,14 Aerococccus urinaehominis11,15 Aerococcus viridansa,5,11 Dolosicoccus16 Dolosigranulum11,17,18 Enterococcusc,d,5 Facklamia languidae,18,19 Facklamia sppe,18,20–22 Gemella haemolysansf,5 Gemella sppf,5,23,24 Globicatella5,25–27 Granulicatella adiacensg,7–9 Granulicatella elegansg,7–9 Helcococcus kunziia,5,28 Ignavigranum18,26,29 Lactococcusd,h,5 Leuconostoc5 Pediococcus5 Streptococcush,i,5 Tetragenococcus5 Vagococcusc,5 Weissella30,31

+ − + − −

+ + + + −

− + + + +

− − + − −

− + V

+ + + + + + + + V

− − + + + + + +